Handbook of Neurochemistry and Molecular Neurobiology: Neurotransmitter Systems [3 ed.] 9780387303512, 9780387303826

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Handbook of Neurochemistry and Molecular Neurobiology Neurotransmitter Systems

Abel Lajtha (Ed.)

Handbook of Neurochemistry and Molecular Neurobiology Neurotransmitter Systems Volume Editor: E. Sylvester Vizi

With 96 Figures and 22 Tables

Editor Abel Lajtha Director Center for Neurochemistry Nathan S. Kline Institute for Psychiatric Research 140 Old Orangeburg Road Orangeburg New York, 10962 USA Volume Editor E. Sylvester Vizi Institute of Experimental Medicine Hungarian Academy of Sciences H-1083 Budapest Szigony 43 Hungary

Library of Congress Control Number: 2006922553 ISBN: 978-0-387-30351-2 Additionally, the whole set will be available upon completion under ISBN: 978-0-387-35443-9 The electronic version of the whole set will be available under ISBN: 978-0-387-30426-7 The print and electronic bundle of the whole set will be available under ISBN: 978-0-387-35478-1 ß 2008 Springer Science+Business Media, LLC. All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Springer Science+Business Media, LLC., 233 Spring Street, New York, NY 10013, USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. springer.com Printed on acid-free paper

SPIN: 11416593 2109 — 5 4 3 2 1 0

Preface

The brain is the organ that collects information from the environment, processes and stores the information, and generates behavior as and when needed. In essence, the brain makes us who we are. For this reason, understanding the biology of brain function is a great challenge and a major goal of modern science. The brain is one of the last great frontiers in science, and the unraveling of its mysteries is comparable in complexity to efforts in space exploration. Therefore, it was not a surprise that the U.S. Congress proclaimed the 1990s the ‘‘Decade of the Brain,’’ a movement also introduced by several other countries, thereby giving a chance for neuroscientists to focus on this topic and to have better conditions for their research. A fundamental goal of neuroscience is to understand how neurons generate behavior and the pathophysiology of different mental and neurological diseases. This requires, among other things, information about where these neurons are located, how they are connected, and how they communicate with each other in various physiological and pathophysiological conditions. At the end of the nineteenth century, the great neuroanatomist Santiago Ramon y Cajal recognized that neurons are the individual signaling elements of the brain. In this book, we focus on these nerve cells of the brain and the chemical molecules that allow them to talk to each other. The discovery of intercellular communication through endogenous molecules is a milestone in the history of science. It makes the brain a unique organ. Our aim is to describe recent discoveries about the basic operations of the brain and to provide an introduction to the adaptations for specific types of information processing. For example, at a chemical synapse, the presynaptic terminal liberates a transmitter substance that acts on the postsynaptic process. Thus, a synapse converts a presynaptic electrical signal into a chemical signal and back into a postsynaptic electrical signal. Current experimental evidence indicates that this assumption needs to be enlarged. Neurons can communicate in a widespread but still organized mode: transmitters released into the extracellular space may have effects on distant extrasynaptic receptors, exerting a tonic effect. Neurons are able to release their transmitters locally into the extracellular space, where the transmitter diffuses slowly over some distance and influences many other neurons. The notion that the amount of transmitter released by an effective action potential arriving at the nerve terminal is not constant but can be modulated by the chemical environment in the vicinity of the release site is now well accepted, as also the effect of transmitters on postsynaptic sites. There is a large body of literature on these subjects, but recent developments are not well covered in textbooks. Therefore, we felt it was time to summarize the available evidence and new ideas on chemical transmission. As part of this goal, shared by our colleagues who agreed to contribute to the book, we asked them to deal with their individual topics so as to produce a balanced review of the current view and to include a discussion of the controversies as well as the accepted facts. We required them also to write for students and interested outsiders, not just for experts in the field. We hope that a reading of this book captures the thinking of cutting-edge developments in neuroscience. This book puts most of its emphasis on different transmitters ranging from acetylcholine to other amines, amino acids, and diffusible gases. By publishing this book, as well as the Handbook of Neurochemistry and Molecular Neurobiology, edited by Dr. Lajtha, in an electronic format, we will be able to provide periodic updates on different topics. We hope the result is appreciated and, more importantly, is helpful and informative to those who wish to learn more about the rapidly growing field of neuroscience.

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To conclude this short introduction, the editor would like to express his gratitude to all who have made this volume possible. It is a pleasure to acknowledge the people who have helped, in a variety ways, with the ˝ to˝, Emese preparation of this book. I thank the authors for their scientific contributions and Ms. Judit Fu Na´na´si, and Dr. Bala´zs Lendvai for their patience and help in the preparation of this volume. E. Sylvester Vizi

Contributors

N. M. Barnes Cellular and Molecular, europharmacology Research Group, Department of Pharmacology, Division of Neuroscience, The Medical School, University of Birmingham, Vincent Drive, Edgbaston, Birmingham B15 2TT, UK B. S. Basavarajappa Division of Analytical Psychopharmacology, New York State Psychiatric Institute, New York, USA; Nathan Kline Institute for Psychiatric Research, Orangeburg, NY, USA; Department of Psychiatry, College of Physicians & Surgeons, Columbia University, New York, NY, USA R. M. Breyer Department of Medicine, Vanderbilt University Medical Center, Nashville, TN, USA M. V. Caldeira Center for Neuroscience and Cell Biology and Department of Zoology, University of Coimbra, 3004, Portugal

T. B. Cooper Division of Analytical Psychopharmacology, New York State Psychiatric Institute, New York, USA; Nathan Kline Institute for Psychiatric Research, Orangeburg, NY, USA; Department of Psychiatry, College of Physicians & Surgeons, Columbia University, New York, NY, USA R. A. Cunha Institute of Biochemistry, Faculty of Medicine, University of Coimbra, 3004–504 Coimbra, Portugal L. Descarries De´partements de pathologie et biologie cellulaire et de physiologie, Centre de recherche en sciences neurologiques, Faculte´ de me´decine, Universite´ de Montre´al, Montre´al, Que´bec, Canada H3C 3J7 C. B. Duarte Center for Neuroscience and Cell Biology and Department of Zoology, University of Coimbra, 3004–517 Coimbra, Portugal

A. L. Carvalho Center for Neuroscience and Cell Biology and Department of Zoology, University of Coimbra, 3004, Portugal

A. C. Dutton Cellular and Molecular, europharmacology Research Group, Department of Pharmacology, Division of Neuroscience, The Medical School, University of Birmingham, Vincent Drive, Edgbaston, Birmingham B15 2TT, UK

A. P. Carvalho Center for Neuroscience and Cell Biology and Department of Zoology, University of Coimbra, 3004, Portugal

I. J. Elenkov San Raffaele Research Center, Laboratory of NeuroEndocrineImmunology, Via dei Bonacolsi snc, Rome 00163, Italy

P. J. Cimino Department of Pathology, University of Washington School of Medicine, Seattle, WA, USA

A. R. Gomes Center for Neuroscience and Cell Biology and Department of Zoology, University of Coimbra, 3004, Portugal

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Contributors B. Gulya´s Department of Clinical Neuroscience, Karolinska Institute, Stockholm, S–171 76 Sweden C. Halldin Department of Clinical Neuroscience, Karolinska Institute, Stockholm, S–171 76 Sweden L. G. Harsing, Jr. Division of Preclinical Research, EGIS Pharmaceuticals Plc, 1165 Budapest, Bokenyfoldi ut 116, Hungary B. L. Hungund Division of Analytical Psychopharmacology, New York State Psychiatric Institute, New York, USA; Nathan Kline Institute for Psychiatric Research, Orangeburg, NY, USA; Department of Psychiatry, College of Physicians & Surgeons, Columbia University, New York, NY, USA

K. S. Montine Department of Pathology, University of Washington School of Medicine, Seattle, WA, USA T. J. Montine Department of Pathology, University of Washington School of Medicine, Seattle, WA, USA S. S. Oja The Centre for Laboratory Medicine, Tampere University Hospital, Tampere, Finland P. Saransaari Brain Research Center, Tampere University Medical School, Tampere, Finland

C. D. Keene Department of Pathology, University of Washington School of Medicine, Seattle, WA, USA

A. Schousboe Department of Pharmacology, Danish University of Pharmaceutical Sciences, DK–2100 Copenhagen, Denmark

J. P. Kiss Laboratory of Drug Resesarch, Department of Pharmacology, Institute of Experimental Medicine, Hungarian Academy of Sciences, H–1083 Budapest, Hungary

B. Sperla´gh Department of Pharmacology, Institute of Experimental Medicine, Hungarian Academy of Sciences, H–1450 Budapest, Hungary

B. Lendvai Department of Pharmacology, Institute of Experimental Medicine, Hungarian Academy of Sciences, H–1083, Szigony u. 43., Budapest, Hungary

M. V. Ugrumov Institute of Developmental Biology RAS, Institute of Normal Physiology RAMS, Moscow, Russia

B. Mazie`re Service Hospitalier Frederic Joliot, Orsay, CEA, F–91403 France N. Mechawar Douglas Hospital Research Centre, McGill University, Verdun, Que´bec, Canada H4H 1R3 R. Meloni Laboratoire de Ge´ne´tique de la Neurotransmission et des Processus Neurode´ne´ne´ratifs. C.N.R.S. UMR 7091, Hoˆpital Pitie´-Salpeˆtrie`re, 75013 Paris, France E. Molna´r MRC Centre for Synaptic Plasticity, Department of Anatomy, University of Bristol, School of Medical Sciences, University Walk, Bristol BS8 1TD, UK

E. S. Vizi Institute of Experimental Medicine, Hungarian Academy of Sciences, H–1083, Szigony u. 43., Budapest, Hungary H. S. Waagepetersen Department of Pharmacology, Danish University of Pharmaceutical Sciences, DK–2100 Copenhagen, Denmark F. Wannenes San Raffaele Research Center, Laboratory of NeuroEndocrineImmunology, Via dei Bonacolsi snc, Rome, 00163, Italy R. Yalamanchili Nathan Kline Institute for Psychiatric Research, Orangeburg, NY, USA

Table of Contents

Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . v Contributors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ix 1

Structural Organization of Monoamine and Acetylcholine Neuron Systems in the Rat CNS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1 L. Descarries . N. Mechawar

2

Brain Neurons Partly Expressing Monoaminergic Phenotype: Distribution, Development, and Functional Significance in Norm and Pathology . . . . . . . . 21 M. V. Ugrumov

3

In Vivo Imaging of Neurotransmitter Systems with PET . . . . . . . . . . . . . . . . . . 75 B. Gulya´s . C. Halldin . B. Mazie`re

4

Synaptic and Nonsynaptic Release of Transmitters . . . . . . . . . . . . . . . . . . . . 101 E. S. Vizi . B. Lendvai

5

Cholinergic Transmission . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 113 B. Lendvai

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Molecular Genetics of Brain Noradrenergic Neurotransmission . . . . . . . . . . . 129 R. Meloni

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Dopamine and the Dopaminergic Systems of the Brain . . . . . . . . . . . . . . . . . 149 L. G. Harsing Jr.

8

5‐Hydroxytryptamine in the Central Nervous System . . . . . . . . . . . . . . . . . . 171 A. C. Dutton . N. M. Barnes

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GABA Neurotransmission: An Overview . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 213 A. Schousboe . H. S. Waagepetersen

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ATP‐Mediated Signaling in the Nervous System . . . . . . . . . . . . . . . . . . . . . . . 227 B. Sperla´gh

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Adenosine Neuromodulation and Neuroprotection . . . . . . . . . . . . . . . . . . . . 255 R. A. Cunha

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Regulation of AMPA Receptors by Metabotropic Receptors and Receptor Tyrosine Kinases: Mechanisms and Physiological Roles . . . . . . . . . . . . . . . . . 275 A. L. Carvalho . M. V. Caldeira . A. R. Gomes . A. P. Carvalho . C. B. Duarte

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Taurine in Neurotransmission . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 325 P. Saransaari . S. S. Oja

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The Endocannabinoid System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 343 B. S. Basavarajappa . R. Yalamanchili . T. B. Cooper . B. L. Hungund

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E Prostanoid Receptors in Brain Physiology and Disease . . . . . . . . . . . . . . . . 385 C. D. Keene . P. J. Cimino . R. M. Breyer . K. S. Montine . T. J. Montine

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Nitric Oxide and other Diffusible Messengers . . . . . . . . . . . . . . . . . . . . . . . . 403 J. P. Kiss

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Molecular Organization and Regulation of Glutamate Receptors in Developing and Adult Mammalian Central Nervous Systems . . . . . . . . . . . . 415 E. Molna´r

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Sympathetic and Peptidergic Innervation: Major Role at the Neural–Immune Interface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 443 I. J. Elenkov . A. Tagliani Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 457

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Structural Organization of Monoamine and Acetylcholine Neuron Systems in the Rat CNS

L. Descarries . N. Mechawar

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Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2

2 2.1 2.2 2.3

Dopamine (DA) Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mesocortical DA System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mesostriatal DA System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Diencephalospinal DA System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

2 3 3 4

3 Noradrenaline (NA) Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 3.1 Coeruleocortical NA System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 3.2 Myelencephalospinal NA System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 4

Adrenaline Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6

5 5.1 5.2 5.3

Serotonin (5-HT) Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Rapheocortical 5‐HT System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Rapheostriatal 5‐HT System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Rapheospinal 5‐HT System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Histamine Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7

7 7.1 7.2 7.3 7.4

Acetylcholine (ACH) Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8 Basalocortical ACh System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8 Septohippocampal ACh System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10 Neostriatal ACh Innervation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10 Spinal ACh Innervation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11

8 8.1 8.2 8.3 8.4

Developmental Aspects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Dopamine Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Noradrenaline Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Serotonin Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acetylcholine Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Concluding Remarks: A New Image of the Neuron . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13

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11 11 12 12 13

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Structural organization of monoamine and acetylcholine neuron systems in the rat CNS

Abstract: The anatomical, cytological, and ultrastructural features of monoamine (dopamine, noradrenaline, adrenaline, serotonin, histamine) and acetylcholine neuron systems have been examined in many regions of mammalian central nervous system, particularly in the rat. By considering these data with an emphasis on innervation densities and ultrastructural relationships, including results obtained during postnatal development, organizational principles and characteristics emerge for each of the modulatory systems. List of Abbreviations: ChAT, choline acetyltransferase; DA, dopamine; DBH, dopamine‐b‐hydroxylase; 5‐HT, 5‐hydroxytryptamine; NA, noradrenaline; PNMT, phenylethanolamine N‐methyltransferase; TH, tyrosine hydroxylase

1

Introduction

Nowadays, neuromodulation may be broadly defined as all actions of neuronally released compounds that produce more than a direct, short‐lived effect on neuronal firing. In this sense, all known neurotransmitters, including the neuropeptides, the purine nucleotides and nucleosides, as well as the gaseous compounds NO and CO, presumably qualify as neuromodulators. In this chapter, however, a more conventional definition is adopted, whereby neurotransmitters exert transient effects through receptors mostly confined to synaptic junctions, whereas neuromodulators may act for longer periods of time through receptors that are mostly located away from release sites. In this view, the major neuromodulatory systems are the catecholamine (dopamine (DA), noradrenaline (NA), and adrenaline), serotonin (5‐HT), histamine, acetylcholine (ACh), and neuropeptide systems, as opposed to the amino acid systems (glutamate, aspartate, glycine, and GABA). Purines, NO, and CO need not be considered as forming systems on their own, since these compounds appear to be always colocalized with modulators and transmitters. Such distinctions are merely operational, however, since it is becoming increasingly clear that one or more of the modulators and/or transmitters are most often coexistent in the same neurons. Owing to their progressive characterization by a variety of chemoanatomical techniques, and particularly fluorescence histochemistry, uptake autoradiography, and immunocytochemistry, the morphological features of the central monoamine and ACh systems can be currently described at three levels of morphological organization: (1) their overall anatomical distribution of constitutive cell bodies of origin and axonal projections throughout the CNS; (2) their regional and intraregional (subnuclear or laminar) distribution of axon terminals (or varicosities) in different territories of innervation; and (3) their ultrastructural characteristics, intrinsic and relational, of putative release sites in the various brain regions. This chapter examines the monoamine and ACh systems of rat brain from this triple standpoint, including data on their development. It focuses mainly on rat, as knowledge in this species is the most complete and detailed. The purpose is not to be exhaustive, but to illustrate some principles as well as organizational features prevailing within and between these systems. In conjunction with the increasing amount of data being currently accrued on the cellular and subcellular distribution of the multiple receptors for each of the neuromodulators, it is thus expected to gain insights into their complementary modes of action and functional properties.

2

Dopamine (DA) Neurons

The general organization of the dopamine (DA) system is rather compartmentalized compared with that of the other monoamines (for a detailed description and bibliographical listing, see Bjo¨rklund and Lindvall, 1984). In the rat CNS, at least six DA projection subsystems have been described—mesostriatal, mesolimbocortical, diencephalospinal, periventricular, incertohypothalamic, and tuberohypophyseal, in addition to DA interneurons in both the olfactory bulb and retina. The mesostriatal and mesolimbocortical DA systems originate from three mesencephalic groups of cell bodies located in the retrorubral field, substantia nigra, and ventral tegmental area, respectively designated as A8, A9, and A10 according to the nomenclature proposed by Dahlstro¨m and Fuxe (1964) at the time of their initial description. The other cell groups giving

Structural organization of monoamine and acetylcholine neuron systems in the rat CNS

1

rise to DA projections, A11 to A15, occupy various parts of the diencephalon. The most caudal, A11, is at the origin of a long descending projection to the spinal cord, whereas the other groups give rise to short projections to the diencephalon and circumscribed hypothalamic and hypophyseal areas. The DA neurons in the olfactory bulb (group A16) are a subset of the periglomerular interneuron population, and thus mostly found in the periglomerular area, with a few also scattered in the outer plexiform layer. In the retina, the A17 group consists of a relatively sparse subpopulation of amacrine cells interspersed in the inner portion of the inner nuclear layer. Thus, the DA systems are constituted by various cell types, from anaxonic to long axon neurons. For a detailed mapping of these cell groups, see Ho¨kfelt et al. (1984b). The number of DA cells in the mesencephalic tegmentum has been estimated at 15,000–20,000 on each side of rat brain (Hedreen and Chalmers, 1972; Guyenet and Crane, 1981; Swanson, 1982): some 9,000 in the ventral tegmental area (Swanson, 1982) and the remainder in the zona compacta of the substantia nigra and retrorubral field. These cells are at the origin of extensive mesotelencephalic projections. Neurons of the A10 group supply the mesolimbocortical system innervating structures such as the amygdala, septum, olfactory tubercle, nucleus accumbens and cerebral cortex, whereas those of the A8 and A9 groups are the main contributors of the nigrostriatal system.

2.1 Mesocortical DA System The DA projection to cerebral cortex is restricted in its distribution, at least in rat. This DA innervation was initially described as confined to the anteromedial or prefrontal, anterior cingulate or pre‐and supragenual, suprarhinal, perirhinal, piriform, and entorhinal cortex (for detailed references to the literature, see Descarries et al., 1987). Additional DA terminal fields were later identified in the dorsomedial frontal area, retrosplenial and adjacent occipital cortex, and in the deep layers of the frontal, parietal, temporal, and occipital neocortex (Descarries et al., 1987). In each of these areas, there is a strong predilection of the DA innervation for certain cortical layers. Moreover, axonal tracing studies have indicated that individual DA neurons innervating the cerebral cortex have fairly circumscribed intracortical territories of projection and do not collateralize extensively (Fallon and Loughlin, 1982; Swanson, 1982; Albanese and Minciacchi, 1983; Loughlin and Fallon, 1984). The heterogeneous distribution of the mesocortical DA subsystem is substantiated by the available data on the number of DA terminals in the different regions of rat cerebral cortex, ranging from 4
104 in layer VI of the occipital cortex to 3.1
106 in layers II–III of the supragenual cingulate cortex (Descarries et al., 1987). Basic neurochemical parameters have been deduced from these numbers. Assuming that cortical DA is mostly concentrated within the varicosities as opposed to intervaricose axon segments, it has been calculated that, depending on the cortical region, the average DA content of a single varicosity should range from 0.6 to 1.7
104 pg of DA for concentrations of 290–810 mg/g or 1.9–5.4
103 M. Interestingly, the figures for mediofrontal and cingulate cortex were very similar to those extrapolated for neostriatal DA varicosities (Doucet et al., 1986). All available observations suggest important regional and perhaps laminar differences in the frequency with which DA axon varicosities in adult rat cerebral cortex make synaptic specializations. Thus, in the anteromedial and the occipital cortex, these axon terminals appear to be mostly if not entirely synaptic (Se´gue´la et al., 1988; Papadopoulos et al., 1989), whereas in the suprarhinal cortex, only 56% display a junctional complex. Relatively low synaptic incidences of 39% and 20% have also been reported for DA terminals in monkey prefrontal and entorhinal cortex, respectively (Smiley and Goldman‐Rakic, 1993; Erickson et al., 2000). In terms of DA function, the significance of such differences between cortical regions has yet to be investigated.

2.2 Mesostriatal DA System Studies of anterogradely labeled axons after injection of biotin dextran in single nigrostriatal neurons have shown that most of these axons travel directly to the striatum, in which they branch abundantly, whereas others branch only sparsely in the striatum and arborize profusely in various extrastriatal structures,

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Structural organization of monoamine and acetylcholine neuron systems in the rat CNS

including globus pallidus and entopeduncular and subthalamic nuclei (Gauthier et al., 1999; see also Prensa and Parent, 2001). In dorsal neostriatum, the density of DA varicosities or terminals, i.e., potential release sites of DA, has been estimated at 1
108/mm3 in the striatal matrix and 1.7
108/mm3 in the striosomes and subcallosal streak (Doucet et al., 1986), which is generally assumed to represent one‐tenth of all terminals in the striatum. Since the volume of rat neostriatum is approximately 45 mm3, the total number of DA varicosities in one striatum must be at least 4.5
109, which represents at least 1660 DA varicosities per neostriatal neuron (for unbiased estimates of neuron number in striatum, see Oorschot, 1996). Then, depending on whether the number of mesencephalic DA neurons projecting to neostriatum is considered to be 3,500 (Ande´n et al., 1966) or 7,000 (Bjo¨rklund and Lindvall, 1984), these individual neurons must be endowed, on average, with 1.3 or 0.6
106 axon varicosities in the striatum, emphasizing the bushy character of their arborization. Furthermore, because only 30%–40% of these DA varicosities are junctional, and 60%–70% do not form synaptic membrane specializations (Descarries et al., 1996), it may be inferred that this DA subsystem operates in large part by diffuse as well as synaptic transmission. Because of the extreme density of this DA innervation, it has also been hypothesized that the spontaneous and evoked release from such a multitude of asynaptic as well as synaptic varicosities might permanently maintain a basal extracellular level of DA throughout the striatum Descarries et al., 1996. This should allow, among other functions and in addition to the transsynaptic effects of DA, for a sustained regulation of widely distributed high‐affinity receptors on neurons, glia, and microvascular elements.

2.3 Diencephalospinal DA System The descending DA projection to the spinal cord originates from a few hundred cells (group A11) located in the dorsal and posterior hypothalamus, paraventricular hypothalamic nucleus, zona incerta, and caudal thalamus (Bjo¨rklund and Skagerberg, 1979; Ho¨kfelt et al., 1979; Swanson et al., 1981). Axons of these DA neurons have been described as bifurcating into an ascending branch to the diencephalon and a descending branch to the spinal cord (Lindvall and Bjo¨rklund, 1974a). In the cord, these descending axons travel partly within lamina I of the dorsal horn and adjoining parts of the dorsal funiculus and partly around the central canal, spreading scattered terminals to the spinal grey at all segmental levels, mainly in laminae III–IV of the dorsal horn, the intermediolateral cell column, the periependymal region, and the ventral horn (Skagerberg et al., 1982; Shirouzu et al., 1990; Ridet et al., 1992). In the intermediolateral cell column and ventral horn, these axon terminals appear to be all synaptic, making junctions with cell bodies and dendrites (Ridet et al., 1992). Around the central canal (at thoracic level), more than two thirds are synaptic, whereas in the dorsal horn, two thirds at the cervical level and one fourth at the thoracic level do not form conventional synapses (Ridet et al., 1992). Thus, as in the cerebral cortex and neostriatum, the diencephalospinal DA system might operate at least partly by diffuse transmission, depending on the region innervated. It is not yet known whether the synaptic and asynaptic DA terminals in the different regions and/or at different levels of the cord arise from collaterals of the same cells.

3

Noradrenaline (NA) Neurons

The entire neuraxis, except for neostriatum, receives a NA innervation, issued from two major clusters of brainstem neurons, one in the locus coeruleus (A6) and its dorsolateral extension (A4) and the other in a series of smaller cell groups occupying the ventrolateral aspect of the medulla and pons (A1, A5, A7) (for a detailed description, see Moore and Card, 1984; see also Ho¨kfelt et al., 1984b). After dopamine‐b‐hydroxylase (DBH) immunostaining, the total number of these cells has been estimated at about 5,000 on each side of the brainstem (Swanson and Hartman, 1975). Noradrenergic neurons are also present in the nucleus tractus solitarii‐dorsal vagal complex and the area postrema (A2). Both the locus coeruleus and lateral tegmental NA groups give rise to ascending and descending projections. The locus coeruleus projects principally to the cerebral cortex, thalamus, cerebellum, and spinal cord, whereas the lateral tegmental groups projects principally to the basal forebrain, hypothalamus, brainstem, and spinal cord (Ungerstedt, 1971; Lindvall and Bjo¨rklund, 1974a, b, 1978).

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3.1 Coeruleocortical NA System The locus coeruleus (A6), comprising approximately 1,500 neuronal cell bodies on each side of the brainstem (Descarries and Saucier, 1972; Swanson, 1976), is at the origin of the entire NA innervation of the cerebral cortex, including hippocampus (Ungerstedt, 1971; for detailed references to the literature, see Audet et al., 1988). A single coeruleocortical axon probably collateralizes from front to back throughout the cortex, infiltrating its whole thickness (Morrison et al., 1981; Nagai et al., 1981; Loughlin et al., 1982). Moreover, studies with retrogradely transported fluorescent dyes have convincingly demonstrated that at least some of the NA neurons innervating the cerebral cortex can concomitantly innervate other distant CNS regions such as the cerebellum or spinal cord (Ade`r et al., 1980; Nagai et al., 1981; Room et al., 1981; Steindler 1981). The intracortical distribution of NA terminals has been studied in considerable detail in rat and monkey. In rat neocortex, the NA axons and their varicosities have been shown to be rather uniformly distributed between the various cytoarchitectonic areas, whereas in primates, there might be a greater degree of regional and laminar heterogeneity (Levitt et al., 1984; Morrison et al., 1984). The average density of NA innervation has been estimated at 1.2
106 varicosities/mm3 of tissue in rat neocortex, with no statistically significant difference between the seven cortical areas examined in the anterior half of the brain (Audet et al., 1988). In every region, the number of NA terminals was greatest in the molecular layer and decreased progressively in the underlying cortex, with a two‐to threefold difference between the upper and lower layers. These numerical data allowed to estimate the possible number of cortical NA varicosities per locus coeruleus cell body of origin (at least 300, 000), their average number per cortical neuron (30–50), their actual incidence among all terminals in the cortex (1/1,000), and the mean endogenous amine content per varicosity (0.22 fg) (Audet et al., 1988). A similar quantitative study in the dorsal hippocampus revealed a more heterogeneous regional and laminar distribution and a significantly higher density of NA innervation, averaging 2.1
106 varicosities/mm3 (Oleskevich et al., 1989). On the basis of a hippocampal volume of 56 mm3, and assuming an equal share of hippocampal NA terminals per neuron, this should represent a further load of 78,400 terminals per locus coeruleus neuron. The number of NA varicosities per hippocampal neuron should range from 20 to 40 per granule cell in the dentate gyrus to 180 per pyramidal cell in CA3, and could represent one varicosity per 880–1,500 synapses, in the DG and CA1, respectively. Similar to neocortex, the NA content per varicosity should be in the order of 0.16–0.21 fg, for concentrations in the 102 M range. In both neocortex and hippocampus, the synaptic incidence of NA varicosities has been shown to be very low, with reported frequencies of 17% or 26% in the parietal cortex (Smiley et al., 1992; Se´gue´la et al., 1990), 7% in the frontal cortex (Cohen et al., 1997), and 16% in the CA1 region of dorsal hippocampus (Umbriaco et al., 1995). A single study in monkey provided a value of 18% for prefrontal cortex (Aoki et al., 1998). Thus, there seems to be a principle of coherence at stake here, whereby a highly divergent projection system such as the coeruleocortical NA system establishes rather loose interrelationships at the ultrastructural level as well, whereas more compartmentalized and more focused projections, such as the mesocortical DA system, will establish more frequent, and perhaps more rigid, synaptic connections (Descarries et al., 1988). Because of its widespread distribution as well as largely asynaptic character, this NA system appears ideally built to act at a distance, on vast neuronal ensembles, and exert rather general, sustained and/or indirect or mediated effects, as expected from a neuromodulator.

3.2 Myelencephalospinal NA System The NA innervation of spinal cord provides further evidence of the extreme divergence of locus coeruleus neurons. Numerous studies have indicated that, although the medullary NA cell groups contribute to this innervation, its principal source is the locus coeruleus (A6) and the A5 and A7 lateral tegmental cell groups (Westlund et al., 1981, 1982, 1983; Fritschy and Grzanna, 1990). The locus coeruleus and adjacent subcoeruleus NA neurons supply input to both the dorsal and ventral horns at all segmental levels, whereas the majority of the NA innervation of the intermediolateral cell column appears to arise from the A5 and A7 groups. The density of this spinal innervation is much greater than that of its DA counterpart. Yet, as

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Structural organization of monoamine and acetylcholine neuron systems in the rat CNS

in the case of the DA system, only 25%–29% of the NA terminals in the dorsal horn would display junctional specializations, whereas a much greater proportion (87% and 85%) is synaptic in the intermediolateral cell column and ventral horn, respectively (Rajaofetra et al., 1992; Ridet et al., 1993).

4

Adrenaline Neurons

Neurons that are immunopositive not only for the biosynthetic enzymes tyrosine hydroxylase (TH) and DBH but also for phenylethanolamine N‐methyltransferase (PNMT), and which presumably synthesize and release adrenaline (Ho¨kfelt et al., 1974), also give rise to long and widely collateralized, albeit less abundant, axonal projections distributed from the forebrain to the spinal cord (for detailed description, see Ho¨kfelt et al., 1984a). The cell bodies of these neurons form three small groups, C1, C2, and C3, confined to medulla oblongata and located near and within the A1 and A2 NA cell groups and on the midline, respectively, within and dorsal to the medial longitudinal fasciculus. PNMT‐immunopositive fibers and nerve endings have been described as concentrated along the ventricular system and most abundant in the bed nucleus of the stria terminalis, various nuclei of hypothalamus, periaqueductal grey matter, brainstem nuclei of visceral afferent and efferent systems, locus coeruleus, and intermediolateral cell column of the spinal cord. In most of these CNS regions, the fine structural features and relationships as well as functional properties of the adrenergic nerve terminals remain to be characterized. In view of its anatomical distribution, this system is assumed to play a significant role in neuroendocrine mechanisms and blood pressure control. In adult rat spinal cord, PNMT‐immunopositive axon terminals have been shown to establish axosomatic and axodendritic synaptic contacts with the preganglionic neurons of the intermediolateral cell column (Milner et al., 1988).

5

Serotonin (5-HT) Neurons

The serotonin (5‐hydroxytryptamine; 5‐HT) system is even more widespread than the NA system, as there is no single region of CNS without a 5‐HT innervation, including each of the circumventricular organs and the cerebroventricular cavity, which is lined by the so‐called supra‐ependymal plexus of varicose 5‐HT fibers (Steinbusch, 1981). Owing to the variety of methodological approaches applicable for the identification and examination of central 5‐HT neurons, this is undoubtedly the transmitter‐defined system about which the most is known in terms of distribution, cytological features, and particularly, ultrastructural relationships. The 5‐HT neuronal cell bodies are mainly found near or in the midline or raphe region of the medulla, pons, and mesencephalon, in nine groups designated B1–B9 according to Dahlstro¨m and Fuxe’s nomenclature (1964). The more caudal groups (B1, B2, and B3 in raphe pallidus, obscurus, and magnus) project mostly to the medulla and spinal cord, whereas the most rostral (B5–B9, in raphe medianus, dorsalis, and the supralemniscal region) provide extensive innervation to the diencephalon and telencephalon. 5‐HT cell bodies have also been found in the area postrema, caudal locus coeruleus, and nucleus interpeduncularis. The nucleus raphe dorsalis is the most prominent, with more than 11,000 5‐HT neurons, representing approximately one‐third of its entire neuron population (Descarries et al., 1982). It is also the best characterized in terms of the cytological features of its constituent neurons and afferent and efferent connectivity. Much as the locus coeruleus NA neurons, it is likely that some of the nucleus raphe dorsalis 5‐HT neurons are highly collateralized and simultaneously project to vast areas of forebrain distant from one another (Fallon and Loughlin, 1982).

5.1 Rapheocortical 5‐HT System The 5‐HT innervations of cerebral cortex, hippocampus, and neostriatum have been the most thoroughly examined. In seven cytoarchitectonic areas from the anterior half of the cerebral cortex, the density of regional and laminar 5‐HT innervation was quantified after uptake labeling of the 5‐HT varicosities in whole hemisphere sections incubated with tritiated 5‐HT in the presence of a monoamine oxidase inhibitor (Audet et al., 1989). The mean regional density of cortical 5‐HT innervation was thus estimated at 5.8
106

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1

varicosities/mm3 of tissue, with significant variations between areas. The highest laminar density was always that of layer I, except in piriform cortex, but each region showed a distinct laminar pattern of 5‐HT innervation. On the basis of these figures, the average number of cortical 5‐HT varicosities per cell body of origin could be calculated to be at least 500,000, their average number per cortical neuron from 145 to 230, their incidence among all cortical axon terminals 1 in 200, and their mean endogenous amine content 0.045 fg for a concentration in the order of 3
103 M. A similar study carried out in the subiculum, Ammon’s horn, and dentate gyrus of the dorsal hippocampus yielded values of 2.7
106 varicosities/mm3 for the average density, but with the values in subiculum > Ammon’s horn > dentate gyrus, and a marked heterogeneity in laminar distribution (Oleskevich and Descarries, 1990). The average number of hippocampal 5‐HT varicosities per cell body of origin could thus be evaluated at 150,000, the number of 5‐HT varicosities per target neuron at 20–130, and the mean endogenous amine content per hippocampal 5‐HT varicosity at 0.05–0.07 fg, a value similar to that in cerebral cortex. In rat neocortex, a detailed study by Se´gue´la and coworkers (1989) has estimated the synaptic incidence of 5‐HT axon varicosities at 36% and 28% in the superficial and deep layers of the frontal cortex, 46% in the parietal cortex, and 37% in the occipital cortex. Even lower frequencies of synaptic specialization were reported for the sensorimotor and prefrontal cortex of monkey (DeFelipe and Jones, 1988; Smiley and Goldman‐Rakic, 1996) and for the cat auditory cortex (DeFelipe et al., 1991). Frequencies of synaptic 5‐HT varicosities ranging between 12% and 24% have also been reported for CA1, CA3, and the dentate gyrus of hippocampus (Oleskevich et al., 1991; Cohen et al., 1995; Umbriaco et al., 1995).

5.2 Rapheostriatal 5‐HT System In neostriatum, the 5‐HT innervation appears rather uniformly distributed, without any suggestion of a patch and matrix pattern. Its density increases from rostral to caudal, however, and is always higher ventrally than dorsally. It ranges from 4.8
106 varicosities/mm3 rostrally to 6.3
106 caudally, for an average of 5.6
106 (Mrini et al., 1995), almost equal to that in cerebral cortex. Such a density corresponds to approximately 90 5‐HT varicosities per neostriatal neuron, thus roughly 18 times less the number for DA terminals. It predicts a value in the order of 0.09 fg for the mean 5‐HT content per neostriatal 5‐HT varicosity, compared to 0.045 fg and 0.06 fg in the cerebral cortex and hippocampus, respectively. The proportion of these 5‐HT varicosities engaged in synaptic contact has been estimated at 10%–13%. In the rostral striatum, for example, this small proportion should correspond to some 4.8
105 synapses/mm3, i.e., approximately 1 in every 2000 striatal synapses.

5.3 Rapheospinal 5‐HT System The 5‐HT innervation of spinal cord is relatively dense, with particular regions of the spinal grey standing out because of their strong 5‐HT innervation: the dorsal horn, particularly lamina I and to a lesser extent lamina II, the ventral horn motor nuclei (laminae VIII and IX), and the intermediolateral cell column in the thoracic cord (Bowker et al., 1982; Skagerberg and Bjo¨rklund, 1985). As in the case of the DA and the NA innervations of the dorsal horn, the 5‐HT innervation in this area has been shown to be only partly synaptic (37%) (Ridet et al., 1993), with little variation between the different laminae of the dorsal horn or at different spinal cord levels (Marlier et al., 1991). In the intermediolateral cell column and anterior horn, however, the 5‐HT varicosities are presumably mostly if not entirely synaptic (Poulat et al., 1992; Ridet et al., 1993), as also reported for DA and NA axon varicosities.

6

Histamine Neurons

Immunocytochemical studies with antibodies against histidine decarboxylase or histamine itself have revealed the existence in the rat brain of a widely distributed network of fine, varicose, unmyelinated axons, originating from small subgroups of nerve cell bodies located in the tuberomammillary nucleus

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Structural organization of monoamine and acetylcholine neuron systems in the rat CNS

(Watanabe et al., 1984; Inagaki et al., 1988; Panula et al., 1989). This system comprises long projections to regions such as the olfactory bulb, cerebral cortex, caudate‐putamen, and thalamus, a relatively dense innervation of numerous areas of hypothalamus, and descending projections to the upper and lower brainstem, cerebellum, and spinal cord. To date, the only light and electron microscopic immunocytochemical description of these nerve fibers and axon varicosities has shown that in cerebral cortex and neostriatum the majority of histamine varicosities do not form synaptic specializations (Takagi et al., 1986).

7

Acetylcholine (ACH) Neurons

Our present knowledge on the structural basis of ACh transmission in the CNS has been largely acquired from the detailed examination of neurons immunostained for choline acetyltransferase (ChAT), the rate‐ limiting enzyme for ACh synthesis. All regions of the CNS are pervaded by dense networks of ChAT‐ immunostained axons originating either from projection neurons located in the basal forebrain or midbrain, and/or from local interneurons (Armstrong et al., 1983; Woolf, 1991). The latter have been shown to contribute either a small fraction (neocortex), almost all (neostriatum), or the totality (spinal cord) of respective regional ACh innervations. Groups of ACh projection neurons and their targets have been extensively described from investigations combining tract‐tracing methods and ChAT immunocytochemistry (Rye et al., 1984; Saper, 1984) and are now commonly referred to as Ch1–Ch8 on the basis of nuclear localization (Mesulam et al., 1983; Mesulam, 1988). The basal forebrain groups Ch1–Ch4 provide for the rich and widespread innervations of neocortex, hippocampus, olfactory bulb, and amygdala. The medial septum and nucleus of the vertical limb of the diagonal band of Broca, Ch1 and Ch2, respectively, send dense ACh projections to the hippocampal formation, while the lateral portion of the horizontal limb of the diagonal band of Broca (group Ch3) innervates mainly the olfactory bulb. Together, groups Ch1–Ch3 also contribute a limited fraction of the total cortical ACh innervation, with axons restricted to limbic areas (cingulate, entorhinal, orbitofrontal and piriform cortex). In contrast, the whole cortical mantle (including these limbic areas) and amygdala receive a rich axon network stemming from the Ch4 neurons in the nucleus basalis magnocellularis of Meynert, which is spread over the substantia innominata and globus pallidus. The pontomesencephalic ACh system (Ch5–Ch6) is the principal projection pathway outside the basal forebrain and has most of its efferents reaching the thalamus and basal forebrain. ACh neurons in the habenula constitute the Ch7 system, which extends projections exclusively to the interpeduncular nucleus via the fasciculus retroflexus pathway. Finally, the parabigeminal nucleus group Ch8, also limited in its scope of innervation, targets the majority of its axons to the superior colliculus (tectum). The availability of a highly sensitive monoclonal antibody with high affinity for whole rat brain ChAT (Cozzari et al., 1990) has allowed the development of experimental conditions leading to the integral staining of ACh axon networks through the full thickness of perfusion‐fixed brain sections. This made it possible to conduct thorough and unbiased electron microscopic descriptions of ChAT‐immunostained axon varicosities (Umbriaco et al., 1994), and led to the development of a semicomputerized light microscopic method to quantify the length of axons and related number of varicosities from ChAT‐ immunostained axon networks (Mechawar et al., 2000). In consequence, our group has produced quantitative descriptions of the cortical, hippocampal, and neostriatal ACh innervations, both in terms of quantified distribution and ultrastructural features (Umbriaco et al., 1994, 1995; Contant et al., 1996; Mechawar et al., 2000; Aznavour et al., 2002). In this section, we mainly discuss these systems (basalocortical, septohippocampal, and neostriatal), as they have been the most thoroughly scrutinized of all central ACh innervations. The ACh innervation intrinsic to the spinal cord will also be described, as it allows for meaningful comparisons with the monoaminergic innervations of this region.

7.1 Basalocortical ACh System It is currently estimated that, in the rat brain, there are 7,000–9,000 Ch4 neurons projecting to the cortex (Rye et al., 1984; Gritti et al., 1993). Although the great majority of ACh axons in cerebral cortex

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originate from the basal forebrain, a 20%–30% portion is contributed by intrinsic bipolar interneurons scattered throughout layers II–VI (Johnston et al., 1981; Eckenstein and Thoenen, 1983; Eckenstein and Baughman, 1984; Levey et al., 1984). In a detailed light microscopic description of ChAT‐immunostained axons distributed in the rat cerebral cortex, 13 different patterns of ACh innervation were identified that generally corresponded to functionally similar cortical areas (Lysakowski et al., 1989). This patterning has recently been associated with modality‐and region‐specific ACh release in neocortex (Fournier et al., 2004), indicating at least a regional control in Ch4 output activity. The laminar and regional densities of the cortical ChAT immunoreactive axon network have been estimated in transverse sections from the frontal (Fr1), parietal (Par1), and occipital (Oc1) cortical areas (Mechawar et al., 2000). The number of varicosities per unit length of axon was counted directly at the microscope and found to be constant throughout these areas (average of 4 varicosities/10 mm of axon). In consequence, the actual number of varicosities in the network could be directly derived from measurements of its length, and the laminar and regional densities of ACh innervation expressed in meters of axon and millions of varicosities per mm3 of tissue. The densest ACh innervation was thus measured in the frontal, followed by the occipital and the parietal cortex, with respective values of 5.4, 4.6, and 3.8
106 varicosities/mm3. In the three areas, layers I and V were the most densely innervated, with respective interareal means of 5.3 and 5.0
106 varicosities/mm3. The least densely innervated were layers IV and VI of the primary sensory areas, with interareal means (Par1 and Oc1) of 3.4 and 3.8
106 varicosities/mm3, respectively. As expected, the laminar distributions were area specific, and characterized by uniformly high densities throughout the frontal cortex and lower densities in layers II/III, IV, and VI of the parietal cortex, as well as in layers IV and VI of the occipital cortex. Compared to monoaminergic innervation densities previously measured in the same areas, the mean density of 4.6
106 ACh varicosities/mm3 for these regions represented the densest of all neuromodulatory inputs to the neocortex. When broken down to individual cells, these figures allow to calculate that, on average, each ACh neuron projecting to the cerebral cortex must be endowed with an axonal arborization totalling at least 0.5 m in length and bearing more than 200,000 varicosities. Moreover, this is likely to be an underestimate, since Ch4 neurons have also been shown to extend several axon collaterals within the basal forebrain, which make synapse onto dendrites of surrounding (unspecified) neurons (Zaborszky and Duque, 2000). This latter observation leads to the conclusion that the activity of basalocortical ACh neurons may itself be subjected to a dual cholinergic modulation: one intrinsic, from its own recurrent axon collaterals; and one extrinsic, from its ACh afferents of the mesopontine tegmentum. The relational features of cortical ACh axon terminals (varicosities) were first described in the primary somatosensory cortex of adult rat (Umbriaco et al., 1994). In all layers of Par1, only a small fraction of these ChAT immunoreactive varicosities were found to form a synaptic contact (junctional complex), i.e., 10%, 14%, 11%, 21%, and 14% in layers I, II/III, IV, V, and VI, respectively, for an interlayer mean of 14%. In general, cortical ACh varicosities were relatively small, and those bearing a synaptic junction were slightly but significantly larger than their nonsynaptic counterparts, i.e., 0.67 versus 0.57 mm in diameter, respectively. The junctional complexes formed by these terminals were single, occupied a small fraction of the total surface of varicosities (3%), and were almost always symmetrical (99%). The relatively few synapses made by ACh varicosities were always axodendritic, either on branches (75%) or on spines (25%). Subsequent investigations in other laboratories have confirmed that the vast majority of ACh varicosities in rat cortex are asynaptic, with reported estimates of 14% and 9% for the frontoparietal and entorhinal region, respectively (Che´dotal et al., 1994; Vaucher and Hamel, 1995). The value of 66% recently reported by Turrini and coworkers (2001) for layer V of rat parietal cortex after labeling with the vesicular ACh transporter was presumably the result of a sampling bias, as suggested by a significantly larger size of the profiles examined in that particular study. In the prefrontal cortex of rhesus monkey, Mrzljak et al. (1995) reported that 44 of 100 serially sectioned ChAT-immunoreactive axon varicosities at the border of layers II and III made synaptic contact, mostly onto small dendritic shafts. In a similar study of two samples of human anterior temporal lobe removed at surgery for epilepsy, Smiley et al. (1997) found 28 of 42 ACh varicosities from layer I and II endowed with small but identifiable synaptic specializations. Whether such variations of synaptic incidence reflect sampling biases, regional differences or species differences remains to be determined. In any event, these

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data allow the conclusion that the actions of ACh depend on both diffuse and synaptic transmission in the neocortex of primates as well as of rat (further discussion in Descarries et al., 2002).

7.2 Septohippocampal ACh System The vast majority of ACh axons in the hippocampus originate from the estimated 7,150 ACh Ch1–Ch2 neurons in each septum (Cadete‐Leite et al., 2003). Indeed, very few bipolar ACh interneurons have been described in the hippocampus, most of which were observed in the stratum lacunosum moleculare of CA1 (Aznavour et al., 2002). As previously found for the neocortical innervation, the periodicity of varicosities along ACh axons remained constant at 4 per 10 mm throughout the dorsal hippocampus, suggesting that this is an intrinsic feature of ACh innervations. The densest laminar ACh innervations were measured in the stratum lacunosum moleculare of CA3, stratum pyramidale of CA1 and CA3, and stratum moleculare of the dentate gyrus, with respective values of 7.5, 8.2, 6.8, and 7.8 million varicosities/mm3 (Aznavour et al., 2002). Regional values were similarly high, ranging from 4.9
106 varicosities/mm3 in CA1 to 6.2
106 varicosities/mm3 in CA3, for an average of 5.9
106 varicosities/mm3 in hippocampus, a value 28% greater than for neocortex and higher than any neuromodulatory input to cortex described so far. Ultrastructural analysis of this innervation in the stratum radiatum of CA1 revealed additional common features with the basalocortical ACh system (Umbriaco et al., 1995). The ACh varicosities in this hippocampal region measured 0.6 mm on average, and only 7% were synaptic. Their few synaptic contacts were symmetrical and made either with dendritic branches or spines. Together with immunoelectron microscopic data demonstrating a predominant extrasynaptic localization of muscarinic ACh receptors in hippocampus or cerebral cortex (reviewed in Volpicelli and Levey, 2004), the above results strongly suggest that the modulatory effects of ACh on cortical function, so well documented in hippocampus, are largely conveyed by diffuse (or volume) transmission in addition to synaptic transmission. In both regions in which this innervation is relatively dense, many of these effects could also depend on the existence of a low ambient level of ACh, permanently maintained in the extracellular space in spite of the presence of acetylcholinesterase (for further discussion, see Descarries et al., 1997).

7.3 Neostriatal ACh Innervation The neostriatum receives by far the densest ACh innervation in the mammalian brain, as manifested by the high measures of cholinergic markers expressed throughout this region. Among others, values of ACh content (Cheney et al., 1975; Hoover et al., 1978), ChAT activity (Hoover et al., 1978), and choline uptake (Rea and Simon, 1981) are particularly elevated. These parameters reflect the profuse ACh axon network originating from cholinergic aspiny interneurons, which are estimated to account for less than 2% of all neostriatal neurons (Woolf and Butcher, 1981; Phelps et al., 1985). ACh interneurons in the caudate and putamen are large cells resembling their basal forebrain counterparts (Armstrong et al., 1983), and also give rise to fine unmyelinated axons periodically adorned with small, round, or ovoid varicosities. Although this local cell population provides the neostriatum with most of its cholinergic innervation, a minor fraction of ACh axons has been found to originate from the Ch5–Ch6 system (Woolf and Butcher, 1981, 1986). The ultrastructural features of these putative release sites were first described by Contant and coworkers (1996), after ChAT‐immunostaining in single thin sections for electron microscopy. As previously found in cerebral cortex and hippocampus, neostriatal ACh varicosities were seldom engaged in synaptic contact. Their frequency of junction of 3% in single sections amounted to 9% when extrapolated to the whole volume of varicosities. The very few ACh synapses were made with synaptic branches (6/10) or spines (4/10). Other axon terminals, unlabeled, were often directly apposed to neostriatal ACh varicosities. Occasional juxtaposition of ChAT‐immunostained varicosities was also observed. From this and a subsequent study in the developing brain (Aznavour et al., 2003), it was concluded that the diffuse mode of transmission was an inherent characteristic of ACh neurons, both as interneurons (neostriatum) and projection neurons (cerebral cortex).

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7.4 Spinal ACh Innervation Unlike the DA, NA, and 5‐HT systems, there are no descending ACh projections from the brain to the spinal cord (Sherriff et al., 1991). Therefore, the spinal ACh innervation originates exclusively from the different categories of spinal ACh neurons, which have been thoroughly described by Barber and collaborators (1984). The preponderant type of ACh neuron in this region is the ventral horn somatic motor neuron, regarded as the canonical ACh neuron since Langley’s seminal work on neuromuscular activation one century ago. The only other ACh neurons with projections leaving the spinal cord are the preganglionic autonomic neurons, located in the intermediate grey matter at thoracolumbar and lumbosacral levels. The three other types of ACh cells in the spinal cord are the small laminae III–V neurons of the dorsal horn, lamina VII partition neurons at the border between the dorsal and ventral horns, and lamina X neurons in the central grey surrounding the central canal. Transverse sections of adult rat spinal cord immunostained for ChAT reveal that somatic motor neurons are organized in central, medial, and lateral motor columns, and that the medial and lateral columns can be further divided into five subcolumns (Barber et al., 1984). There are apparently widespread intra‐ and intercolumnar interactions, as suggested by extent of intermingling longitudinal and transverse motor dendrite bundles. Likewise, at autonomic levels, the somata and dendritic arborizations of partition cells and central canal ACh neurons are heavily mixed with those of autonomic neurons. These extensive interconnections between morphologically diverse ACh neurons have been described as the basis of a cholinergic propriospinal system (Sherriff and Henderson, 1994; Huang et al., 2000). A rich ACh axon network pervades all layers and regions of the spinal cord, including the ependymal cell layer (Phelps et al., 1984; Scha¨fer et al., 1998). These varicose axons originate in various (undetermined) proportions from the different types of ACh neurons mentioned above. Initial reports have shown ChAT‐ immunoreactive terminals to contact both ACh and non‐ACh elements in the spinal cord. The current prevalent view is that these terminals are mostly, if not exclusively, synaptic in nature. This view is likely biased by the fact that most studies on the subject have focused specifically on contacts made on the cell bodies and proximal dendrites of motoneurons, Renshaw cells, and sympathetic preganglionic neurons (Markham and Vaughn, 1990; Alvarez et al., 1999). Of these descriptions, the ones concerning the ACh innervation on motoneuron somatodendrites are of particular interest. This innervation, which is thought to arise from the canal cluster cells, consists of C‐type cholinergic terminals characterized by periodic postsynaptic specializations called subsurface cisterns (Nagy et al., 1993).

8

Developmental Aspects

The early development of monoamine and ACh neurons in mammalian CNS has led to the notion that these modulatory systems play significant roles in the installment and refinement of neuronal connectivity during brain maturation. In rat, for example, dopaminergic fibers enter the cortical plate just before birth (Kalsbeek et al., 1988); noradrenergic fibers reach the anterior parietal cortex around E20, and occipital cortex at birth (Verney et al., 1982); serotoninergic fibers and terminals (Seiger and Olson, 1973), as well as pioneering cholinergic fibers (Mechawar and Descarries, 2001), are already seen in the cortical plate at birth (for reviews, see Semba, 1992, 2004). Yet, only two laboratories have examined the morphological features of these systems at electron as well as light microscopic levels during development: the University of Thessaloniki group, in Greece, which has focused on the monoamine systems, and our own laboratory, in Montreal, which has mainly studied the developing cholinergic system.

8.1 Dopamine Neurons The ultrastructural features of DA neurons have been examined in two brain regions during development: lateral septum and striatum. The DA innervation of the lateral septum was reported to undergo a marked reorganization during the first two postnatal weeks, when it acquired features comparable to the adult (Antonopoulos et al., 1997). The ultrastructural analysis suggested that there might be two different DA inputs to this region: the first developing earlier in life, affecting remote parts of neurons through

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symmetrical axodendritic synapses, and the second developing later and affecting neuronal somata through asymmetrical axosomatic synapses. In the striatum, the same authors described the DA innervation of caudate‐putamen and nucleus accumbens as exhibiting a similar type of synaptic connectivity throughout development, but evolving from symmetrical synapses mostly located on dendritic shafts at an early stage to a later stage where symmetrical axospinous synapses also became a prominent feature (Antonopoulos et al., 2002). Interestingly, this study also indicated that after initial rises to values >90% and 80% in the islands and matrix of caudate‐putamen, respectively, and almost 100% and 75% in the shell and core of the nucleus accumbens, the proportion of DA varicosities making synapses declined after P7 in the caudate‐putamen and after P14 in the nucleus accumbens to respective values of 63% and 35% in the islands and matrix of caudate‐ putamen, and 71% and 47% in the shell and core of nucleus accumbens.

8.2 Noradrenaline Neurons The noradrenergic innervations of the septum, motor and visual cortex, and dorsal lateral geniculate nucleus were described in developing rat brain, by means of light and electron microscopic immunocytochemistry with DBH antibodies (Latsari et al., 2002, 2004; Antonopoulos et al., 2004). In all four regions, few, relatively thick NA fibers were present at birth, which arborized gradually into an adult pattern of thinner varicose fibers by the second postnatal week, and reached the adult density of innervation one week later. Irrespective of the postnatal age examined, only a minority of these NA varicosities displayed synaptic specializations (10%–15%), which were usually symmetrical and found on dendritic branches. It was concluded from these data that, in these brain regions at least, transmission by diffusion is the major mode of NA action in the developing as well as adult brain.

8.3 Serotonin Neurons The growth of 5‐HT innervations was examined in numerous subcortical regions of postnatal rat brain: lateral septum (Dinopoulos et al., 1993), lateral ventricles (Dinopoulos and Dori, 1995), lateral geniculate (Dinopoulos et al., 1995), basal forebrain (Dinopoulos et al., 1997), superior colliculus, and ventrolateral thalamic nucleus (Dori et al., 1998). Acquisition of such data from the developing cortex (Dori et al., 1996) was complicated by the fact that, within the first weeks after birth, thalamocortical neurons transiently express 5‐HT plasma membrane transporter and vesicular monoamine transporter (Lebrand et al., 1996) and are thus immunoreactive for 5‐HT and indistinguishable from bona fide 5‐HT neurons. A constant feature of the 5‐HT innervations in the postnatal period, albeit parenchymal or intraventricular, was their progressive change from a few, thick and smooth unmyelinated axonal fibers at birth to a ramified and relatively dense network of fine varicose axons, infiltrating the whole region and reaching its adult pattern of distribution and density within the first three weeks. At the ultrastructural level, however, much more diversity was apparent. In the lateral septum, 5‐HT varicosities were described as almost always synaptic and showing symmetrical synapses, whether on somata, dendritic shafts, or spines. In the dorsal portion of the lateral septum, they formed characteristic pericellular basket‐like arrangements around cell somata and their primary dendrites, as previously described for DA terminals (Descarries and Beaudet, 1983). In the lateral ventricles, the 5‐HT varicosities were located close to the ventricular surface of the ependymal lining, but never made synapses on ependymal cells, even when morphologically mature (shape, size, and content). In the lateral geniculate and basal forebrain, the synaptic frequency of the 5‐HT varicosities displayed a biphasic temporal profile. The proportion of varicosities forming synapses was reported to increase from birth to the end of the second postnatal week, and then decline markedly in the third week before increasing again to adult values of about 40% in both regions. A similar biphasic pattern was also described in the superficial layers of the superior colliculus (involved in visual functions); whereas in its deep layers and in the ventrolateral thalamic nucleus (areas involved in motor functions), the proportion of 5‐HT varicosities engaged in synaptic contact showed a continuous increase from birth to adulthood, to become a fully synaptic innervation making mostly symmetrical synapses onto dendritic shafts.

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8.4 Acetylcholine Neurons A recent description of the ACh innervation in the frontal, parietal, and occipital neocortex of the postnatal rat has revealed that this ACh system develops rapidly and much earlier than previously suspected (Mechawar and Descarries, 2001). At birth, a few ChAT‐immunostained fibers capped with growth cones are already seen throughout the cortical plate and marginal zone. By P4, faintly immunoreactive interneurons are first detected. Rapid ingrowth and proliferation ensues resulting in an adult‐like distribution of a highly elaborate network of fine varicose ACh axons by P8. In the parietal area, adult densities of innervation are reached by P16, while development continues until the end of the first month in the frontal area and even later in the occipital area. Two parameters of this ingrowth have been quantified (Mechawar and Descarries, 2001): the elongation (and branching) of ACh axons and their number of axon varicosities per unit length. This latter value was shown to increase steadily in all cortical layers and areas, doubling from 2 varicosities per 10 mm of axon at P4 to the adult ratio of 4 per 10 mm at P16. It was thus possible to calculate that, within the first two weeks after birth, a single basalocortical neuron produced an average of 2 cm of axon and 9,000 varicosities/day, i.e., almost 1 mm of axon and 400 varicosities/hour. A similar study has also been carried out in CA1, CA3, and the dentate gyrus of dorsal hippocampus (Aznavour et al., 2005). At P8, an elaborate network of varicose ChAT‐immunostained axons was already present in all three hippocampal regions. As in neocortex, the number of axon varicosities per unit length of ACh axon increased during the first two weeks after birth to reach the adult value of 4 per 10 mm at P16. At this age, the laminar distribution of this network resembled that of maturity, but adult densities of axons and axon varicosities were reached only by P32. Between P8 and P32, the mean densities in the three regions increased from 8.4 to 14 m of axons, and 2.3 to 5.7 million varicosities/mm3 of tissue. This suggested that, on average, the septohippocampal ACh neurons are capable of generating 7.5 cm of axon and 27,000 varicosities/day, i.e., more than 3 mm of axon and 1,120 varicosities/hour. Such growth rates are even higher than previously estimated for nucleus basalis ACh neurons innervating the neocortex, and emphasize the remarkable growth capacities of ACh neurons. These studies have also examined the intrinsic and relational features of ACh varicosities in the developing neocortex and hippocampus (Mechawar et al., 2002; Aznavour et al., 2005). In both regions, these varicosities were of similar size throughout development and only slightly smaller than in the adult. They were endowed with aggregated synaptic vesicles, and the frequency with which they showed a mitochondrion increased gradually with age, from about 20% at P8 to >40% at P32. As in the adult, the vast majority of these varicosities were asynaptic throughout the postnatal period. Since the proportion of synaptic ACh varicosities was stable during development, it could be inferred that the number of ACh synapses had already reached its adult value by the end of the second week, at least in the parietal cortex (0.55
106/mm3 at P16, compared to 0.53
106 at >P60). The postnatal growth and ultrastructural characteristics of a developing ACh innervation have also been examined in the neostriatum, where this innervation arises almost completely from a limited number of large interneurons. As in cortex and hippocampus, the invasion of neostriatum by ACh axons mostly takes place during the first two weeks after birth, but continues at a slower rate until the fourth postnatal week (Aznavour et al., 2003), in keeping with biochemical measurements of ChAT activity (Coyle and Yamamura, 1976). As in neocortex and hippocampus, the intrinsic and relational ultrastructural features of these rapidly growing axons were examined at three developmental time points, i.e., after the first, second, and fourth postnatal weeks (P8, P16, and P32) (Aznavour et al., 2003). Again, a low synaptic incidence was measured at all three ages, indicating that this structural feature is an intrinsic determinant of the functioning of this system during development as well as in the adult.

9

Concluding Remarks: A New Image of the Neuron

The detailed comparison of the structural attributes of neuromodulatory systems, whether at the level of their general organization, regional distribution, or fine structure, underscores some consistent features of the major modulatory systems. First and foremost, it requires a drastic revision of what may be called the

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traditional (textbook) image of the neuron, mostly inherited from the study of the neuromuscular junction. Thus, relatively dense and widespread innervations by fine, unmyelinated axons whose multiple branches bear innumerable varicosities (terminals) appears to be the rule for most if not all modulatory neurons in the adult and developing CNS. It may also be assumed that these axonal arborizations, endowed with varicosities that often lack a junctional specialization, are highly plastic, not only in a functional sense but also in their structural configuration. As initially postulated by Descarries and coworkers (1975, 1977; Beaudet and Descarries, 1978), it is likely that such small varicosities, lying free in the neuropil, undergo incessant movements of remodeling and translocation along their parent fibers, resulting in release of their transmitter or modulator in different microlocations of the tissue at different moments in time. Several lines of evidence suggest that many of the peptidergic systems might also share such properties. It is also apparent that these structural determinants apply to both extrinsic and intrinsic neuromodulatory systems, as defined by Katz and Frost (1996). By definition, extrinsic neuromodulation refers to the capacity of a system to cause global changes, affecting many functional circuits simultaneously. The best examples of extrinsic modulatory systems are the DA, NA, 5‐HT, and almost all ACh systems. To varying degrees, these systems pervade most CNS regions and originate from relatively small numbers of neuronal cell bodies, grouped in discrete nuclei. Thus, through their innervation of numerous cortical and subcortical regions, the DA subsystems contribute to various aspects of motor function, neuroendocrine control, motivation, and behavioral learning. The NA, 5‐HT, and ACh systems are even more widespread, innervating most if not all regions of the CNS. They seem to be involved in capacities of a rather global nature and rely on the simultaneous functioning of widely distributed numerous neuronal circuits, such as waking and sleep, arousal, attention, emotional states, learning, memory, and ultimately consciousness. In this perspective, state‐ and context‐dependent responsiveness of the nervous system may be viewed as resulting from the joint activity of multiple, spatially and temporally overlapping extrinsic neuromodulatory systems, as well as that of more specialized, function‐specific systems. Intrinsic neuromodulation arises from neurons entirely contained within a given circuitry (i.e., interneurons). The massive ACh innervation of neostriatum issued from a fraction of its relatively small population of scattered interneurons is a good example of an intrinsic modulatory system. These tonically active neurons appear to be critical elements in the striatal circuitry controlling motor planning, movement, and associative learning (Graybiel et al., 1994; Bennett and Wilson, 1999). Theoretically, intrinsic neuromodulation produces local changes in neuronal computation within a circuit. In view of the largely asynaptic nature of the neostriatal ACh system, its influence is certainly exerted beyond point‐to‐point synaptic connections, and presumably reaches a variety of more‐or‐less distant cellular targets within the neostriatal circuitry. As an intrinsic component, the level of activity of this ACh system would be directly dependent on that of the overall circuitry, the latter being itself modulated by extrinsic inputs (e.g., DA and 5‐HT). It is tempting to speculate that the activity of such a circuitry is reflected by fluctuations in the ambient level of its intrinsic neuromodulators (see Descarries et al., 1997). On the other hand, the ambient levels of extrinsic neuromodulators should more closely depend on the activity of these systems themselves. Much as the progressive unraveling of the numerous modulatory systems, several fairly recent data and concepts pertaining to chemical neurotransmission per se emphasize the previously unsuspected multiplicity of transmission modes in the CNS. The coexistence of transmitter substances within the same neurons (Ho¨kfelt et al., 1980; Merighi, 2002) is now recognized as a common if not universal trait in the CNS, much as the release of transmitter from dendrites, and presumably from cell bodies (Cheramy et al., 1981; He´ry et al., 1982). Spillover beyond the synaptic clefts has been demonstrated for most of the highly if not exclusively synaptic, amino acid transmitters (Isaacson et al., 1993; Kullmann, 2000). There is also ample evidence that multiple metabotropic and/or ionotropic receptors exist for every neurotransmitter/modulator, and receptors for a wide variety of transmitters, and even different receptor subtypes for the same transmitter, are expressed by a given neuron. Moreover, whenever visualized by electron microscopic immunocytochemistry, most neuronal receptors, whether somatodendritic and/or axonal, appear to be located on extra‐ as well as intrasynaptic portions of the plasma membrane (e.g., Riad et al., 2000). Lastly, a growing number of signaling cascades, accounting for short‐, medium‐, and long‐term effects, have been identified for most

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neurotransmitter/modulators. All these mechanisms and properties are obviously needed to support the coordinated activity of a complex nervous system, carrying out multiple functions simultaneously and capable of processing highly diversified information and to react, adapt, and learn from it.

References Ade`r JP, Room P, Postema F, Korf J. 1980. Bilaterally diverging axon collaterals and contralateral projections from rat locus coeruleus neurons, demonstrated by fluorescent retrograde double labeling and norepinephrine metabolism. J Neural Transm 49: 207-218. Albanese A, Minciacchi D. 1983. Organization of the ascending projection from the ventral tegmental area. A multiple fluorescent retrograde tracer study in the rat. J Comp Neurol 216: 406-420. Alvarez FJ, Dewey DE, McMillin P, Fyffe REW. 1999. Distribution of cholinergic contacts on Renshaw cells in the rat spinal cord: A light microscopic study J Physiol 515: 787-797. Ande´n NE, Fuxe K, Hamberger B, Ho¨kfelt T. 1966. A quantitative study of the nigro‐striatal dopamine neuron system in the rat. Acta Physiol Scand 67: 306-312. Antonopoulos J, Dinopoulos A, Dori I, Parnavelas JG. 1997. Distribution and synaptology of dopaminergic fibers in the mature and developing lateral septum of the rat. Brain Res Dev Brain Res 102: 135-141. Antonopoulos J, Dori I, Dinopoulos A, Chiotelli M, Parnavelas JG. 2002. Postnatal development of the dopaminergic system of the striatum in the rat. Neuroscience 110: 245-256. Antonopoulos J, Latsari M, Dori I, Chiotelli M, Parnavelas JG, et al. 2004. Noradrenergic innervation of the developing and mature septal area of the rat. J Comp Neurol 476: 80-90. Aoki C, Venkatesan C, Go CG, Forman R, Kurose H. 1998. Cellular and subcellular sites for noradrenergic action in the monkey dorsolateral prefrontal cortex as revealed by the immunocytochemical localization of noradrenergic receptors and axons. Cereb Cortex 8: 269-277. Armstrong DM, Saper CB, Levey AI, Wainer BH, Terry RD. 1983. Distribution of cholinergic neurons in rat brain: Demonstrated by the immunocytochemical localization of choline acetyltransferase. J Comp Neurol 216: 53-68. Audet MA, Descarries L, Doucet G. 1989. Quantified regional and laminar distribution of the serotonin innervation in the anterior half of adult rat cerebral cortex. J Chem Neuroanat 2: 29-44. Audet MA, Doucet G, Oleskevich S, Descarries L. 1988. Quantified regional and laminar distribution of the noradrenaline innervation in the anterior half of the adult rat cerebral cortex. J Comp Neurol 274: 307-318.

Aznavour N, Mechawar N, Descarries L. 2002. Comparative analysis of the cholinergic innervation in the dorsal hippocampus of adult mouse and rat. A quantitative immunocytochemical study. Hippocampus 12: 406-417. Aznavour N, Mechawar N, Watkins KC, Descarries L. 2003. Fine structural features of the acetylcholine innervation in the developing neostriatum of rat. J Comp Neurol 460: 280-291. Aznavour N, Watkins KC, Descarries L. 2005. Postnatal development of the cholinergic innervation in the dorsal hippocampus of rat: A quantitative light and electron microscopic immunocytochemical description. J Comp Neurol 486: 61-75. Barber RP, Phelps PE, Houser CR, Crawford GD, Salvaterra PM, et al. 1984. The morphology and distribution of neurons containing choline acetyltransferase in the adult rat spinal cord: An immunocytochemical study. J Comp Neurol 229: 329-346. Beaudet A, Descarries L. 1978. The monoamine innervation of rat cerebral cortex: Synaptic and nonsynaptic relationships. Neuroscience 3: 851-860. Bennett BD, Wilson CJ. 1999. Spontaneous activity of neostriatal cholinergic interneurons in vitro. J Neurosci 19: 5586-5596. Bjo¨rklund A, Lindvall O. 1984. Dopamine‐containing systems in the CNS. Handbook of Chemical Neuroanatomy, Vol. 2. Classical Transmitters in the CNS. Bjo¨rklund A, Ho¨kfelt T, editors. Amsterdam: Elsevier; pp. 55-122. Bjo¨rklund A, Skagerberg G. 1979. Evidence for a major spinal cord projection from the diencephalic A11 dopamine cell group in the rat using transmitter‐specific fluorescent retrograde tracing. Brain Res 177: 170-175. Bowker RM, Westlund KN, Sullivan MC, Coulter JD. 1982. Organization of descending serotonergic projections to the spinal cord. Prog Brain Res 57: 239-298. Cadete‐Leite A, Pereira PA, Madeira MD, Paula‐Barbosa MM. 2003. Nerve growth factor prevents cell death and induces hypertrophy of basal forebrain cholinergic neurons in rats withdrawn from prolonged ethanol intake. Neuroscience 119: 1055-1069. Che´dotal A, Umbriaco D, Descarries L, Hartman BK, Hamel E. 1994. Light and electron microscopic immunocytochemical analysis of the neurovascular relationships of choline acetyltransferase and vasoactive intestinal polypeptide nerve terminals in the rat cerebral cortex. J Comp Neurol 343: 57-71.

15

16

1

Structural organization of monoamine and acetylcholine neuron systems in the rat CNS

Cheney DL, Le Fevre HF, Racagni G. 1975. Choline acetyltransferase activity and mass fragmentographic measurement of acetylcholine in specific nuclei and tracts of rat brain. Neuropharmacology 14: 801-809. Cheramy A, Leviel V, Glowinski J. 1981. Dendritic release of dopamine in the substantia nigra. Nature 289: 537-542. Cohen Z, Ehret M, Maıˆtre M, Hamel E. 1995. Ultrastructural analysis of tryptophane hydroxylase immunoreactive nerve terminals in the rat cerebral cortex and hippocampus: Their associations with local blood vessels. Neuroscience 66: 555-569. Cohen Z, Molinatti G, Hamel E. 1997. Astroglial and vascular interactions of noradrenaline terminals in the rat cerebral cortex. J Cereb Blood Flow Metab 17: 894-904. Contant C, Umbriaco D, Garcia S, Watkins KC, Descarries LD. 1996. Ultrastructural characterization of the acetylcholine innervation in adult rat neostriatum. Neuroscience 71: 937-947. Coyle JT, Yamamura HI. 1976. Neurochemical aspects of the ontogenesis of cholinergic neurons in the rat brain. Brain Res 118: 429-440. Cozzari C, Howard J, Hartman B. 1990. Analysis of epitopes on choline acetyltransferase (ChAT) using monoclonal antibodies (Mabs). Soc Neurosci Abstr 16: 200. Dahlstro¨m A, Fuxe K. 1964. Evidence for the existence of monoamine‐containing neurons in the central nervous system I. Demonstration of monoamines in the cell bodies of brain stem neurons. Acta Physiol Scand 62 (Suppl. 232): 1-55. DeFelipe J, Jones EG. 1988. A light and electron microscopic study of serotonin‐immunoreactive fibers and terminals in the monkey sensory‐motor cortex. Exp Brain Res 71: 171-182. DeFelipe J, Hendry SHC, Hashikawa T, Jones EG. 1991. Synaptic relationships of serotonin‐immunoreactive terminal baskets of GABA neurons in the cat auditory cortex. Cereb Cortex 1: 117-133. Descarries L, Beaudet A. 1983. The use of radioautography for investigating transmitter‐specific neurons. Handbook of Chemical Neuroanatomy, Vol. 1, Methods in Chemical Neuroanatomy. Bjo¨rklund A, Ho¨kfelt T, editors. Amsterdam: Elsevier; pp. 286-364. Descarries L, Saucier G. 1972. Disappearance of the locus coeruleus in the rat after intraventricular 6‐hydroxydopamine. Brain Res 37: 310-316. Descarries L, Beaudet A, Watkins KC. 1975. Serotonin nerve terminals in adult rat neocortex. Brain Res 100: 563-588. Descarries L, Doucet G, Lemay B, Se´gue´la P, Watkins KC. 1988. Structural basis of cortical monoamine function. Neurotransmitters and Cortical Function. Avoli M, Reader TA, Dykes RW, Gloor P, editors. New York: Plenum Publishing, pp. 321-332.

Descarries L, Gisiger V, Steriade M. 1997. Diffuse transmission by acetylcholine in the CNS. Prog Neurobiol 53: 603-625. Descarries L, Lemay B, Doucet G, Berger B. 1987. Regional and laminar density of the dopamine innervation in adult rat cerebral cortex. Neuroscience 21: 807-824. Descarries L, Mechawar N, Aznavour N. 2002. Structural determinants of the roles of acetylcholine in cerebral cortex. Progress in Brain Research, Vol. 145, Acetylcholine in the Cerebral Cortex. Descarries L, Krnjevic K, Steriade M, editors. Amsterdam: Elsevier; pp. 45-58. Descarries L, Watkins KC, Garcia S, Beaudet A. 1982. The serotonin neurons in nucleus raphe dorsalis of adult rat. A light and electron microscope radioautographic study. J Comp Neurol 207: 239-254. Descarries L, Watkins KC, Garcia S, Bosler O, Doucet G. 1996. Dual character, asynaptic and synaptic, of the dopamine innervation in adult rat neostriatum: A quantitative autoradiographic and immunocytochemical analysis. J Comp Neurol 375: 167-186. Descarries L, Watkins KC, Lapierre Y. 1977. Noradrenergic axon terminals in the cerebral cortex of rat. III. Topometric ultrastructural analysis. Brain Res 133: 197-222. Dinopoulos A, Dori I. 1995. The development of the serotonergic fiber network of the lateral ventricles of the rat brain: A light and electron microscopic immunocytochemical analysis. Exp Neurol 133: 73-84. Dinopoulos A, Dori I, Parnavelas JG. 1993. Serotonergic innervation of the mature and developing lateral septum of the rat: A light and electron microscopic immunocytochemical analysis. Neuroscience 55: 209-222. Dinopoulos A, Dori I, Parnavelas JG. 1995. Serotonergic innervation of the lateral geniculate nucleus of the rat during postnatal development: A light and electron microscopic immunocytochemical analysis. J Comp Neurol 363: 532-544. Dinopoulos A, Dori I, Parnavelas JG. 1997. The serotonin innervation of the basal forebrain shows a transient phase during development. Brain Res Dev Brain Res 99: 38-52. Dori I, Dinopoulos A, Blue ME, Parnavelas JG. 1996. Regional differences in the ontogeny of the serotonergic projection to the cerebral cortex. Exp Neurol 138: 1-14. Dori IE, Dinopoulos A, Parnavelas JG. 1998. The development of the synaptic organization of the serotonergic system differs in brain areas with different functions. Exp Neurol 154: 113-125. Doucet G, Descarries L, Garcia S. 1986. Quantification of the dopamine innervation in adult rat neostriatum. Neuroscience 19: 427-445. Eckenstein F, Baughman RW. 1984. Two types of cholinergic innervation in cortex, one co‐localized with vasoactive intestinal polypeptide. Nature 309: 153-155.

Structural organization of monoamine and acetylcholine neuron systems in the rat CNS Eckenstein F, Thoenen H. 1983. Cholinergic neurons in the rat cerebral cortex demonstrated by immunohistochemical localization of choline acetyltransferase. Neurosci Lett 36: 211-215. Erickson SL, Sesack SR, Lewis DA. 2000. The dopamine innervation of monkey entorhinal cortex: Postsynaptic targets of tyrosine hydroxylase terminals. Synapse 36: 47-56. Fallon JH, Loughlin SE. 1982. Monoamine innervation of the forebrain: Collateralization. Brain Res Bull 9: 295-307. Fournier GN, Semba K, Rasmusson DD. 2004. Modality‐and region‐specific acetylcholine release in the rat neocortex. Neuroscience 126: 257-262. Fritschy JM, Grzanna R. 1990. Demonstration of two separate descending noradrenergic pathways to the rat spinal cord: Evidence for an intragriseal trajectory of locus coeruleus axons in the superficial layers of the dorsal horn. J Comp Neurol 291: 553-582. Gauthier J, Parent M, Le´vesque M, Parent A. 1999. The axonal arborization of single nigrostriatal neurons in rats. Brain Res 834: 228-232. Graybiel AM, Aosaki T, Flaherty AW, Kimura M. 1994. The basal ganglia and adaptive motor control. Science 265: 1826-1831. Gritti I, Mainville L, Jones BE. 1993. Codistribution of GABA‐ with acetylcholine‐synthesizing neurons in the basal forebrain of the rat. J Comp Neurol 329: 438-457. Guyenet PG, Crane JK. 1981. Non‐dopaminergic nigro‐striatal pathway. Brain Res 213: 291-305. Hedreen JC, Chalmers JP. 1972. Neuronal degeneration in rat brain induced by 6‐hydroxydopamine: A histological and biochemical study. Brain Res 47: 1-36. He´ry F, Faudon M, Ternaux JP. 1982. In vivo release of serotonin in two raphe nuclei (raphe dorsalis and magnus) of the cat. Brain Res Bull 8: 123-129. Ho¨kfelt T, Fuxe K, Goldtein M, Johansson O. 1974. Evidence for the existence of adrenaline neurons in the rat brain. Brain Res 66: 235-251. Ho¨kfelt T, Johansson O, Goldstein M. 1984a. Central catecholamine neurons as revealed by immunohistochemistry with special reference to adrenaline neurons. Handbook of Chemical Neuroanatomy, Vol. 2, Classical Transmitters in the CNS. Bjo¨rklund A, Ho¨kfelt T, editors. Amsterdam: Elsevier; pp. 157-276. Ho¨kfelt T, Ma˚rtensson, Bjo¨rklund A, Kleinau S, Goldstein M. 1984b. Distributional maps of tyrosine‐hydroxylase‐immunoreactive neurons in the rat brain. Handbook of Chemical Neuroanatomy, Vol. 2, Classical Transmitters in the CNS. Bjo¨rklund A, Ho¨kfelt T, editors. Amsterdam: Elsevier; pp. 277-379. Ho¨kfelt T, Lundberg JM, Schultzberg M, Johansson O, Ljungdahl A, et al. 1980. Coexistence of peptides and

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putative transmitters in neurons. Adv Biochem Psychopharmacol 22: 1-23. Ho¨kfelt T, Phillipson O, Goldtein M. 1979. Evidence for a dopaminergic pathway in the rat descending from the A11 cell group to the spinal cord. Acta Physiol Scand 107: 393-395. Hoover DB, Muth EA, Jacobowitz DM. 1978. A mapping of the distribution of acetylcholine, choline acetyltransferase and acetylcholinesterase in discrete areas of rat brain. Brain Res 153: 295-306. Huang A, Noga BR, Carr PA, Fedirchuk B, Jordan LM. 2000. Spinal cholinergic neurons activated during locomotion: Localization and electrophysiological characterization. J Neurophysiol 83: 3537-3547. Inagaki N, Yamatodani A, o‐Yamamoto M, Tohyama M, Watanabe T, et al. 1988. Organization of histaminergic fibers in the rat brain. J Comp Neurol 273: 283-300. Isaacson JS, Solis JM, Nicoll RA. 1993. Local and diffuse synaptic actions of GABA in the hippocampus. Neuron 10: 165-175. Johnston MV, McKinney M, Coyle JT. 1981. Neocortical cholinergic innervation: A description of extrinsic and intrinsic components in the rat. Exp Brain Res 43: 159-172. Kalsbeek A, Voorn P, Buijs RM, Pool CW, Uylings HB. 1988. Development of the dopaminergic innervation in the prefrontal cortex of the rat. J Comp Neurol 269: 58-72. Katz PS, Frost WN. 1996. Intrinsic neuromodulation: Altering neuronal circuits from within. Trends Neurosci 19: 54-61. Kullmann DM. 2000. Spillover and synaptic cross talk mediated by glutamate and GABA in the mammalian brain. Prog Brain Res 125: 339-351. Latsari M, Antonopoulos J, Dori I, Chiotelli M, Dinopoulos A. 2004. Postnatal development of the noradrenergic system in the dorsal lateral geniculate nucleus of the rat. Brain Res Dev Brain Res 149: 79-83. Latsari M, Dori I, Antonopoulos J, Chiotelli M, Dinopoulos A. 2002. Noradrenergic innervation of the developing and mature visual and motor cortex of the rat brain: A light and electron microscopic immunocytochemical analysis. J Comp Neurol 445: 145-158. Lebrand C, Cases O, Adelbrecht C, Doye A, Alvarez C, et al. 1996. Transient uptake and storage of serotonin in developing thalamic neurons. Neuron 17: 823-835. Levey AI, Wainer BH, Rye DB, Mufson EJ, Mesulam MM. 1984. Choline acetyltransferase‐immunoreactive neurons intrinsic to rodent cortex and distinction from acetylcholinesterase neurons. Neuroscience 13: 341-353. Levitt P, Rakic P, Goldman‐Rakic P. 1984. Comparative assessment of monoamine afferents in mammalian cerebral cortex. Monoamine Innervation of Cerebral Cortex. Descarries L, Reader TA, Jasper HH, editors. New York: Alan R. Liss; pp. 41-59.

17

18

1

Structural organization of monoamine and acetylcholine neuron systems in the rat CNS

Lindvall O, Bjo¨rklund A. 1974a. The glyoxylic acid fluorescence histochemical method: A detailed account of the methodology for the visualization of central catecholamine neurons. Histochemistry 39: 97-127. Lindvall O, Bjo¨rklund A. 1974b. The organization of the ascending catecholamine neuron systems in the rat brain as revealed by the glyoxylic acid fluorescence method. Acta Physiol Scand (Suppl. 412): 1‐48. Lindvall O, Bjo¨rklund A. 1978. Organization of catecholamine neurons in rat central nervous system. Handbook of Psychopharmacology, Vol. 9. Iversen L, Iversen S, Snyder SN, editors. New York: Plenum Press; pp. 139-231. Loughlin SE, Fallon JH. 1984. Substantia nigra and ventral tegmental area projections to cortex: Topography and collateralization. Neuroscience 11: 425-435. Loughlin SE, Foote SL, Fallon SH. 1982. Locus coeruleus projections to cortex: Topography, morphology and collateralization. Brain Res Bull 9: 287-294. Lysakowski A, Wainer BH, Bruce G, Hersh LB. 1989. An atlas of the regional and laminar distribution of choline acetyltransferase immunoreactivity in rat cerebral cortex. Neuroscience 28: 291-336. Markham JA, Vaughn JE. 1990. Ultrastructural analysis of choline acetyltransferase‐immunoreactive sympathetic preganglionic neurons and their dendritic bundles in rat thoracic spinal cord. Synapse 5: 299-312. Marlier L, Sandillon F, Poulat P, Rajaofetra, Geffard M, Privat A. 1991. Serotonergic innervation of the dorsal horn of rat spinal cord: Light and electron microscopic immunocytochemical study. J Neurocytol 20: 310-322. Mechawar N, Descarries L. 2001. The cholinergic innervation develops early and rapidly in the rat cerebral cortex: A quantitative immunocytochemical study. Neuroscience 108: 555-567. Mechawar N, Cozzari C, Descarries L. 2000. Cholinergic innervation of adult rat cerebral cortex: A quantitative immunocytochemical description. J Comp Neurol 428: 305-318. Mechawar N, Watkins KC, Descarries L. 2002. Ultrastructural features of the acetylcholine innervation in the developing parietal cortex of rat. Neuroscience 111: 83-94. Merighi A. 2002. Costorage and coexistence of neuropeptides in the mammalian CNS. Prog Neurobiol 66: 161-190. Mesulam MM. 1988. Central cholinergic pathways: Neuroanatomy and some behavioral implications. Neurotransmitters and Cortical Function: From Molecules to Mind. Avoli M, Reader TA, Gloor D, Dykes RW, editors. New York: Plenum Press; pp. 237-260. Mesulam MM, Mufson EJ, Wainer BH, Levey AI. 1983. Central cholinergic pathways in the rat: An overview based on an alternative nomenclature (Ch1–Ch6). Neuroscience 10: 1185-1201.

Milner TA, Morrison SF, Abate C, Reis DJ. 1988. Phenylethanolamine N‐methyltransferase‐containing terminals synapse directly on sympathetic preganglionic neurons in the rat. Brain Res 17: 205-222. Moore RY, Card JP. 1984. Noradrenaline‐containing neuron systems. Handbook of Chemical Neuroanatomy, Vol. 2, Classical Transmitters in the CNS. Bjo¨rklund A, Ho¨kfelt T, editors. Amsterdam: Elsevier; pp. 123-156. Morrison JH, Foote SL, Bloom FE. 1984. Regional, laminar, developmental, functional characteristics of noradrenaline and serotonin innervation patterns in monkey cortex. Monoamine Innervation of Cerebral Cortex. Descarries L, Reader TA, Jasper HH, editors. New York: Alan R. Liss; pp. 61-75. Morrison JH, Molliver ME, Grzanna R, Coyle JT. 1981. The intracortical trajectory of the coeruleo‐cortical projection in the rat: A tangentially organized cortical afferent. Neuroscience 6: 139-18. Mrini A, Soucy JP, Lafaille F, Lemoine P, Descarries L. 1995. Quantification of the serotonin hyperinnervation in adult rat neostriatum after neonatal 6‐hydroxydopamine lesion of nigral dopamine neurons. Brain Res 669: 303-308. Mrzljak L, Pappy M, Leranth C, Goldman-Rakic PS. 1995. Cholinergic synaptic circuitry in the macaque prefrontal cortex. J Comp Neurol 357: 603-617. Nagai T, Satoh K, Imamoto K, Maeda T. 1981. Divergent projections of catecholamine neurons of the locus coeruleus as revealed by fluorescent retrograde double labeling techniques. Neurosci Lett 23: 117-123. Nagy JI, Yamamoto T, Jordan LM. 1993. Evidence for the cholinergic nature of c‐terminals associated with subsurface cisterns in a‐motoneurons of rat. Synapse 15: 17-32. Oleskevich S, Descarries L. 1990. Quantified distribution of the serotonin innervation in adult rat hippocampus. Neuroscience 34: 19-33. Oleskevich S, Descarries L, Lacaille JC. 1989. Quantified distribution of the noradrenaline innervation in the hippocampus of adult rat. J Neurosci 9: 3803-3815. Oleskevich S, Descarries L, Watkins KC, Se´gue´la P, Daszuta A. 1991. Ultrastructural features of the serotonin innervation in adult rat hippocampus. An immunocytochemical description in single and serial thin sections. Neuroscience 42: 777-791. Oorschot DE. 1996. Total number of neurons in the neostriatal, pallidal, subthalamic, substantia nigral nuclei of the rat basal ganglia: A stereological study using the Cavalieri and optical dissector methods. J Comp Neurol 366: 580-599. Panula P, Pirvola U, Auvinen S, Airaksinen MS. 1989. Histamine immunoreactive nerve fibers in the rat brain. Neuroscience 28: 585-610. Papadopoulos GC, Parnavelas JG, Buijs RM. 1989. Light and electron microscopic immunocytochemical analysis of the

Structural organization of monoamine and acetylcholine neuron systems in the rat CNS dopamine innervation of the rat visual cortex. J Neurocytol 18: 303-310. Phelps PE, Barber RP, Houser CR, Crawford GD, Salvaterra PM, et al. 1984. Postnatal development of neurons containing choline acetyltransferase in rat spinal cord: An immunocytochemical study. J Comp Neurol 229: 347-361. Phelps PE, Houser CR, Vaughn JE. 1985. Immunocytochemical localization of choline acetyltransferase within the rat neostriatum: A correlated light and electron microscopic study of cholinergic neurons and synapses. J Comp Neurol 238: 286-307. Poulat P, Marlier L, Rajaofetra N, Privat A. 1992. 5‐Hydroxytryptamine, substance P and thyrotropin‐releasing hormone synapses in the intermediolateral cell column of the rat thoracic spinal cord. Neurosci Lett 136: 19-22. Prensa L, Parent A. 2001. The nigrostriatal pathway in the rat: A single‐axon study of the relationship between dorsal and ventral tier nigral neurons and the striosome/matrix striatal compartments. J Neurosci 21: 7247-7260. Rajaofetra N, Ridet JL, Poulat P, Marlier L, Sandillon F, et al. 1992. Immunocytochemical mapping of noradrenergic projections to the rat spinal cord with an antiserum against noradrenaline. J Neurocytol 21: 481-494. Rea MA, Simon JR. 1981. Regional distribution of cholinergic parameters within the rat striatum. Brain Res 219: 317-326. Riad M, Garcia S, Watkins KC, Jodoin N, Doucet E´, et al. 2000. Somatodendritic localization of 5‐HT1A and preterminal axonal localization of 5‐HT1B serotonin receptors in adult rat brain. J Comp Neurol 417: 181-194. Ridet JL, Rajaofetra N, Teilhac JR, Geffard M, Privat A. 1993. Evidence for non‐synaptic serotonergic and noradrenergic innervation of the rat dorsal horn and possible involvement of neuron‐glia interactions. Neuroscience 52: 143-157. Ridet JL, Sandillon F, Rajaofetra N, Geffard M, Privat A. 1992. Spinal dopaminergic system of the rat: Light and electron microscopic study using an antiserum against dopamine, with particular emphasis on synaptic incidence. Brain Res 598: 233-241. Room P, Postema F, Korf J. 1981. Divergent axon collaterals of rat locus coeruleus. Brain Res 221: 219-230. Rye DB, Wainer BH, Mesulam MM, Mufson EJ, Saper CB. 1984. Cortical projections arising from the basal forebrain: A study of cholinergic and noncholinergic components employing combined retrograde tracing and immunohistochemical localization of choline acetyltransferase. Neuroscience 13: 627-643. Saper CB. 1984. Organization of cerebral cortical afferent systems in the rat. II. Magnocellular basal nucleus. J Comp Neurol 222: 313-342. Scha¨fer MK, Eiden LE, Weihe E. 1998. Cholinergic neurons and terminal fields revealed by immunohistochemistry for

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the vesicular acetylcholine transporter. I. Central nervous system. Neuroscience 84: 331-359. Se´gue´la P, Watkins KC, Descarries, L. 1988. Ultrastructural features of dopamine axon terminals in the anteromedial and suprarhinal cortex of adult rat. Brain Res 442: 11-22. Se´gue´la P, Watkins KC, Descarries L. 1989. Ultrastructural relationships of serotonin axon terminals in the cerebral cortex of the adult rat. J Comp Neurol 289: 129-142. Se´gue´la P, Watkins KC, Geffard M, Descarries L. 1990. Noradrenaline axon terminals in adult rat neocortex: An immunocytochemical analysis in serial thin sections. Neuroscience 35: 249-264. Seiger A, Olson L. 1973. Late prenatal ontogeny of central monoamine neurons in the rat: Fluorescence histochemical observations. Z Anat Entwicklungsgesch 140: 281-318. Semba K. 1992. Development of central cholinergic neurons. Handbook of Chemical Neuroanatomy, Vol. 10, Ontogeny of Transmitters and Peptides in the CNS. Bjo¨rklund A, Ho¨kfelt T, Tohyama M, editors. Amsterdam: Elsevier; pp. 33-62. Semba K. 2004. Phylogenetic and ontogenetic aspects of the basal forebrain cholinergic neurons and their innervation of the cerebral cortex. Descarries L, Krnjevic K, Steriade M, editors. Progress in Brain Research, Vol. 145, Acetylcholine in the Cerebral Cortex. Amsterdam: Elsevier; pp. 3-43. Sherriff FE, Henderson Z. 1994. A cholinergic propriospinal innervation of the rat spinal cord. Brain Res 634: 150-154. Sherriff FE, Henderson ZH, Morrison JFB. 1991. Further evidence for the absence of a descending cholinergic projection from the brainstem to the spinal cord in the rat. Neurosci Lett 128: 52-56. Shirouzu M, Anraku T, Iwashi TA, Yoshida M. 1990. A new dopaminergic terminal plexus in the ventral horn of the spinal cord. Immunohistochemical studies at the light and electron microscopic levels. Experientia 46: 201-204. Skagerberg G, Bjo¨rklund A. 1985. Topographic principles in the spinal projections of serotonergic and non‐ serotonergic brainstem neurons in the rat. Neuroscience 15: 445-480. Skagerberg G, Bjo¨rklund A, Lindvall O, Schmidt RH. 1982. Origin and termination of the diencephalo‐spinal dopamine system in the rat. Brain Res Bull 9: 237-244. Smiley JF, Goldman‐Rakic PS. 1993. Heterogeneous targets of dopamine synapses in monkey prefrontal cortex demonstrated by serial section electron microscopy: A laminar analysis using the silver‐enhanced diaminobenzidine sulfide (SEDS) immunolabeling technique. Cereb Cortex 3: 223-238. Smiley JF, Goldman‐Rakic PS. 1996. Serotonergic axons in monkey prefrontal cerebral cortex synapse predominantly on interneurons as demonstrated by serial section electron microscopy. J Comp Neurol 367: 431-443.

19

20

1

Structural organization of monoamine and acetylcholine neuron systems in the rat CNS

Smiley JF, Morrell F, Mesulam MM. 1997. Cholinergic synapses in human cerebral cortex: An ultrastructural study in serial sections. Exp Neurol 144: 361-368. Smiley JF, Williams SM, Szigeti K, Goldman‐Rakic PS. 1992. Light and electron microscopic characterization of dopamine‐immunoreactive axons in human cerebral cortex. J Comp Neurol 32: 325-335. Steinbusch HWM. 1981. Distribution of serotonin in the central nervous system of the rat—cell bodies and terminals. Neuroscience 6: 557-618. Steindler DA. 1981. Locus coeruleus neurons have axon collaterals that branch to the forebrain and cerebellum. Brain Res 223: 367-373. Swanson LW. 1976. The locus coeruleus: A cytoarchitectonic Golgi and immunohistochemical study in the albino rat. Brain Res 110: 39-56. Swanson LW. 1982. The projections of the ventral tegmental area and adjacent regions: A combined fluorescent retrograde tracer and immunofluorescence study in the rat. Brain Res Bull 9: 321-353. Swanson LW, Hartman BK. 1975. The central adrenergic system: An immunofluorescence study of the location of cell bodies and their efferent connections in the rat utilizing dopamine‐b‐hydroxylase as a marker. J Comp Neurol 163: 467-506. Swanson LW, Sawchenko PE, Be´rod A, Hartman BK, Helle KB, et al. 1981. An immunohistochemical study of the organization of catecholaminergic cells and terminal fields in the paraventricular and supraoptic nuclei of the hypothalamus. J Comp Neurol 196: 271-285. Takagi H, Morishima Y, Matsuyama T, Hayashi H, Watanabe T, et al. 1986. Histaminergic axons in the neostriatum and cerebral cortex of the rat: A correlated light and electron microscopic immunocytochemical study using histidine decarboxylase as a marker. Brain Res 364: 155-168. Turrini P, Casu MA, Wong TP, De Koninck Y, Ribeiro‐Da‐ Silva A, et al. 2001. Cholinergic nerve terminals establish classical synapses in the rat cerebral cortex: Synaptic pattern and age‐related atrophy. Neuroscience 105: 277-285. Umbriaco D, Garcia S, Beaulieu C, Descarries L. 1995. Relational features of acetylcholine, noradrenaline, serotonin and GABA axon terminals in the stratum radiatum of adult rat hippocampus (CA1). Hippocampus 5: 605-620. Umbriaco D, Watkins KC, Descarries L, Cozzari C, Hartman BK. 1994. Ultrastructural and morphometric features of the acetylcholine innervation in adult rat parietal cortex: An electron microscopic study in serial sections. J Comp Neurol 348: 351-373.

Ungerstedt U. 1971. Stereotaxic mapping of the monoamine pathways in the rat brain. Acta Physiol Scand 367: (Suppl) 1-48. Vaucher E, Hamel E. 1995. Cholinergic basal forebrain neurons project to cortical microvessels in the rat: Electron microscopic study with anterogradely transported Phaseolus vulgaris leucoagglutinin and choline acetyltransferase immunocytochemistry. J Neurosci 15: 7427-7441. Verney C, Berger B, Adrien J, Vigny A, Gay M. 1982. Development of the dopaminergic innervation of the rat cerebral cortex. A light microscopic immunocytochemical study using anti-tyrosine hydroxylase antibodies. Brain Res 281: 41-52. Volpicelli LA, Levey AI. 2004. Muscarinic acetylcholine receptor subtypes in cerebral cortex and hippocampus. Prog Brain Res 145: 59-66. Watanabe T, Taguchi Y, Shiosaka S, Tanaka J, Kubota H, et al. 1984. Distribution of histaminergic neuron system in the central nervous system of rats; a fluorescent immunocytochemical analysis with histidine decarboxylase as a marker. Brain Res 295: 13-25. Westlund KN, Bowker RM, Ziegler MG, Coulter JD. 1981. Origins of spinal noradrenergic pathways demonstrated by retrograde transport of antibody to dopamine‐b‐hydroxylase. Neurosci Lett 25: 243-249. Westlund KN, Bowker RM, Ziegler MG, Coulter JD. 1982. Descending noradrenergic projections and their spinal terminations. Prog Brain Res 57: 219-238. Westlund KN, Bowker RM, Ziegler MG, Coulter JD. 1983. Noradrenergic projections to the spinal cord of the rat. Brain Res 263: 15-31. Woolf NJ. 1991. Cholinergic systems in mammalian brain and spinal cord. Prog Neurobiol 37: 475-524. Woolf NJ, Butcher LL. 1981. Cholinergic neurons in the caudate‐putamen complex proper are intrinsically organized: A combined Evans blue and acetylcholinesterase analysis. Brain Res Bull 7: 487-507. Woolf NJ, Butcher LL. 1986. Cholinergic systems in the rat brain. III. Projections from the pontomesencephalic tegmentum to the thalamus, tectum, basal ganglia, basal forebrain. Brain Res Bull 16: 603-637. Zaborszky L, Duque A. 2000. Local synaptic connections of basal forebrain neurons. Behav Brain Res 115: 143-158. Zoli M, Jansson A, Sykova´ E, Agnati LF, Fuxe K. 1999. Volume transmission in the CNS and its relevance for neuropsychopharmacology. Trends Pharmacol Sci 20: 142-150.

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Brain Neurons Partly Expressing Monoaminergic Phenotype: Distribution, Development, and Functional Significance in Norm and Pathology

M. V. Ugrumov

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Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 23

2 2.1 2.1.1 2.1.2 2.1.3 2.2 2.3

Brain Neurons Partly Expressing the MA‐ergic Phenotype in Adult Mammals . . . . . . . . . . . . . . . Neurons Expressing Individual Enzymes of MA Synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Hypothalamus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Striatum . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Other Brain Regions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Bienzymatic TH- and AADC-Expressing Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Non-MA-ergic Neurons Expressing the MA Transporters . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

26 26 26 29 31 32 33

3 3.1 3.1.1 3.1.2 3.2

Brain Neurons Partly Expressing MA‐ergic Phenotype in Mammals in Ontogenesis . . . . . . . . . Neurons Expressing Individual Enzymes of MA Synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Hypothalamus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Extrahypothalamic Regions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Non-MA-ergic Neurons Expressing the MA Transporters . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

35 35 35 39 42

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Functional Properties and Functional Significance of the Neurons Partly Expressing the MA‐ergic Phenotype . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Monoenzymatic Neurons Expressing TH . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Monoenzymatic Neurons Expressing AADC . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ensembles of Monoenzymatic Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Non‐MA‐ergic Neurons Expressing the MA Transporters . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

44 44 47 48 51

4.1 4.2 4.3 4.4 5

Tuberoinfundibular Neurons Partly Expressing DA‐ergic Phenotype in Hyperprolactinemia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 52

6 6.1 6.2 6.3 6.4

Striatal Neurons Partly Expressing DA‐ergic Phenotype in Parkinson’s Disease . . . . . . . . . . . . . . Monoenzymatic TH‐Expressing Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Monoenzymatic AADC‐Expressing Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Bienzymatic TH‐ and AADC‐Expressing Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Origin, Functional Properties, and Functional Significance of Striatal Neurons Partly or Completely Expressing the DA‐ergic Phenotype . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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2008 Springer ScienceþBusiness Media, LLC.

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Brain neurons partly expressing monoaminergic phenotype

Regulation of the Partial Expression of MA‐ergic Phenotype by the Brain Neurons in Norm and Pathology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Regulation of the Partial Expression of MA‐ergic Phenotype by Neural Afferents . . . . . . . . . . . . . . Paracrine Regulation of the Partial Expression of MA‐ergic Phenotype by Diffusive Factors . . Hormonal Regulation of the Partial Expression of MA‐ergic Phenotype . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract: In addition to the monoaminergic (MA‐ergic) neurons possessing the whole set of enzymes of monoamine (MA) synthesis from the precursor amino acid and the MA membrane transporter, the neurons partly expressing the MA‐ergic phenotype have been first discovered almost twenty years ago. Most of the neurons expressing individual enzymes of MA synthesis lack the MA transporter. These so‐ called monoenzymatic neurons are widely distributed throughout the brain in adult mammals being even more numerous than MA‐ergic neurons. Individual enzymes of MA synthesis are expressed continuously or transiently over certain periods of ontogenesis and in adulthood under functional insufficiency of the MA‐ergic neurons, e.g., under their chronic stimulation or in certain neurodegenerative diseases. The earlier data suggest an important functional role of monoenzymatic neurons. Most monoenzymatic neurons possess enzymes of dopamine (DA) synthesis, tyrosine hydroxylase (TH), or aromatic L‐amino acid decarboxylase (AADC). TH and AADC are enzymatically active in a substantial number of monoenzymatic neurons being capable to convert L‐tyrosine to L‐3,4‐dihydroxyphenylalanine (L‐DOPA) and L‐DOPA to DA or serotonin, respectively. L‐DOPA produced in monoenzymatic TH‐neurons is supposed to play a role of a neurotransmitter or a neuromodulator providing its action on the target neurons via catecholamine receptors. Moreover, L‐DOPA released from the monoenzymatic TH‐neurons is captured by monoenzymatic AADC‐neurons or dopaminergic (DA‐ergic) and serotoninergic neurons for DA synthesis (Kannari et al., 2006). Such cooperative synthesis of MAs is considered as a compensatory reaction under the failure of MA‐ergic neurons, e.g., in neurodegenerative diseases like hyperprolactinemia and Parkinson’s disease which are developed primarily because of the degeneration of DA‐ergic neurons of the tuberoinfundibular system and the nigrostriatal system, respectively. Noteworthy, the neurotoxin‐induced increased level of prolactin returns with time to the normal level due to stimulation of DA synthesis by the neurons of the tuberoinfundibular system, most probably because of the turning on cooperative synthesis of DA by monoenzymatic neurons. The same compensatory mechanism is supposed to be used under the failure of the nigrostriatal DA‐ergic system that is manifested by the increased number of monoenzymatic neurons in the striatum of animals with neurotoxin‐induced parkinsonism and in humans with Parkinson’s disease. Expression of the enzymes of MA synthesis in non‐MA‐ergic neurons is controlled by intercellular signals such as classical neurotransmitters (catecholamines), neurotrophic factors (brain‐derived neurotrophic factor, glia‐derived neurotrophic factor), and perhaps hormones (prolactin, estrogens, progesterone). Thus, a substantial number of the brain neurons express partly the MA‐ergic phenotype, mostly individual complementary enzymes of MA synthesis, serving to produce MAs in cooperation that is considered as a compensatory reaction under the failure of MA‐ergic neurons. List of Abbreviations: AADC, aromatic L‐amino acid decarboxylase; AN, arcuate nucleus; DA, dopamine; DOPA, 3,4‐dihydroxyphenylalanine; GABA, g‐aminobutyric acid; GTP, guanosine triphosphate; MA(s), monoamine(s); MPTP, 1‐methyl‐4‐phenyl‐1,2,3,6‐tetrahydropyridine; 6‐OHDA, 6‐hydroxydopamine; SCN, suprachiasmatic nucleus; TH, tyrosine hydroxylase; VMAT2, vesicular monoamine transporter 2

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Introduction

Mechanisms of the regulation of the brain development, plasticity, and integration via chemical neurotransmission are the crucial issue of Neuroscience. The neural and neuroendocrine regulations of most important functions are under the control of dozens or even hundreds of neurotransmitters, neuromodulators, and neurohormones. Monoamines (MAs), catecholamines (dopamine (DA), noradrenaline, and adrenaline), and serotonin are among the most important classical neurotransmitters which are widely distributed all over the brain in ontogenesis and adulthood (Squire et al., 2003). The most frequent MAs, DA and serotonin, are produced enzymatically from L‐tyrosine and L‐tryptophan (the precursor amino acids) in the cytosol of both cell bodies and processes (> Figures 2‐1 and > 2‐2). L‐tyrosine is converted first to L‐3,4‐dihydroxyphenylalanine (L‐DOPA) by tyrosine hydroxylase (TH), the rate‐limiting enzyme of catecholamine synthesis, and then to DA by aromatic L‐amino acid decarboxylase (AADC). In turn, L‐tryptophan is transformed first to 5‐hydroxytryptophan with tryptophan hydroxylase, the rate limiting enzyme of serotonin synthesis, and then to serotonin by AADC

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. Figure 2‐1 Schematic representation of the functioning of dopaminergic and serotoninergic neurons. A, precursor amino acid, L‐tyrosine or L‐tryptophan; D, aromatic L‐amino acid decarboxylase; M, monoamine, dopamine or serotonin; T, tyrosine hydroxylase or tryptophan hydroxylase; TC, target cell; X, intermediate synthetic product, L‐3, 4‐dihydroxyphenylalanine or 5‐hydroxytryptophan, , dopamine or serotonin transporter; , receptor to dopamine or serotonin; , secretory granule at high magnification; , vesicular monoamine transporter 2

. Figure 2‐2 Synthetic pathways of serotonin (a) and dopamine (b)

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(> Figure 2‐2). Noteworthy, the enzymatic activity of AADC greatly exceeds that of TH and tryptophan hydroxylase (Moore et al., 1985). DA and serotonin, the final synthetic products in DA‐ergic and serotoninergic neurons, are captured from cytosol to the secretory granules (¼dense core vesicles) by the vesicular membrane transporter 2 (VMAT2) (> Figure 2-1) (Hoffman et al., 1998; Weihe and Eiden, 2000). In noradrenergic and adrenergic neurons, DA is subsequently converted into noradrenaline and adrenaline by intragranular enzymes, DA b‐hydroxylase and phenyletanolamine N‐methyltransferase, respectively (Squire et al., 2003). MAs stored in secretory granules are discharged via exocytosis. After the action on the targets and partial enzymatic degradation, MAs are captured by the MA‐specific membrane transporter into the monoaminergic (MA‐ergic) neurons for subsequent reutilization (> Figure 2-1) (Hoffman et al., 1998). Although MAs act on the neurons in ontogenesis and adulthood via the same specific receptors their final effects are quite different. They provide a long‐lasting irreversible (¼morphogenetic, imprinting) action on the differentiating target neurons regulating the expression of their specific phenotype in ontogenesis and a short‐term reversible action on the differentiated target neurons controlling their functional activity in adulthood (Ugrumov, 1997). In the early sixties, the MA‐ergic neurons containing catecholamines and serotonin have been first detected on sections with the histofluorescent technique (Dahlstro¨m and Fuxe, 1964) that gave an opportunity to map the catecholaminergic and serotoninergic neurons in the brain. Later, this mapping has been confirmed and slightly modified by using additional morphological approaches; autoradiography of the neurons following the administration of the radiolabeled MAs (Beaudet and Descarries, 1979), and immunostaining of the enzymes of MA synthesis (Ho¨kfelt et al., 1984) or MAs (Steinbusch, 1984). The pioneer morphological studies of the Swedish and Dutch groups were followed by an accumulation of physiological evidence that the clusters of MA‐ergic neurons or the so‐called MA‐ergic centers located in a number of brain regions are involved in neural and neuroendocrine regulations of central and visceral functions, e.g., general metabolism, water–mineral metabolism, memory, different types of behavior, circadian rhythms, etc. (Montange and Calas, 1988; Weiner et al., 1988). Although the MA‐ergic neurons have been already studied for about 40 years, there is a discrepancy in their definition. Indeed, until presently, the MA‐ergic neurons are identified on sections by detecting individual specific markers or an incomplete set of specific markers: (1) MAs (mono‐immunolabeling) (Bjo¨rklund and Lindvall, 1984; Steinbusch, 1984); (2) individual, most often first rate‐limiting enzymes of MA synthesis (mono‐immunolabeling) (Ho¨kfelt et al., 1984); (3) the whole set of the enzymes of MA synthesis (double‐ and triple‐labeling) (Ikemoto et al., 1999); (4) MA membrane transporters (mono‐ and double labeling) (Lorang et al., 1994; Hoffman et al., 1998); (5) colocalization of individual enzymes of MA synthesis and MAs (double‐immunolabeling) (Ershov et al., 2002a, b); (6) colocalization of individual enzymes of MA synthesis and the MA transporter (double‐immunolabeling and/or a combination of immunocytochemistry with in situ hybridization) (Lorang et al., 1994; Betarbet et al., 1997; Cossette et al., 2005a); (7) VMAT2 and its colocalization with the earlier markers (Hoffman et al., 1998; Weihe et al., 2006). Based on the data accumulated over the last 40 years, one may define the MA‐ergic neurons as those having specific machinery providing the MA turnover, i.e., the neurons expressing the MA transporter, the whole set of the enzymes of MA synthesis from the precursor amino acid, and VMAT2. However, VMAT2 is a semi‐specific marker being common for all MA‐ergic neurons, catecholaminergic, and serotoninergic neurons (Hoffman et al., 1998; Weihe and Eiden, 2000). The neurons partly expressing MA‐ergic phenotype have been first discovered in the eighties when the brain regions containing the TH‐immunoreactive (IR) neurons but lacking AADC‐immunoreactive neurons, and vice versa were detected (Jaeger et al., 1983, 1984; Okamura et al., 1988b). The existence of the so‐called monoenzymatic neurons was definitely proved by using double‐immunolabeling of the enzymes of DA and serotonin synthesis (Meister et al., 1988; Ikemoto et al., 1998a, b; Ershov et al., 2002a, b). Another type of neurons partly expressing MA‐ergic phenotype possess the MA transporter but lack either the whole set of the enzymes of MA synthesis or only the first rate‐limiting enzyme (De Vitry et al., 1986; Ugrumov et al., 1989a; Hoffman et al., 1998). The number of neurons partly expressing the MA‐ergic phenotype in ontogenesis and in adulthood of mammals with neurodegenerative diseases (hyperprolactinemia and Parkinson’s disease) appears to exceed those in normal adult mammals (Meredith et al., 1999; Porritt et al., 2000; Ershov et al., 2002a).

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Brain neurons partly expressing monoaminergic phenotype

The discovery of neurons partly expressing the MA‐ergic phenotype has raised a number of crucial questions: (1) what is their functional significance; (2) whether they are numerous and widely distributed over the brain; (3) if the conventional classification of MA‐ergic neurons based on histofluorescence of MAs (Dahlstro¨m and Fuxe, 1964; Bjo¨rklund and Lindvall, 1984) or immunostaining of the first rate‐limiting enzymes of MA synthesis (Ho¨kfelt et al., 1984) or MAs (Steinbusch, 1984) is still valid. The answer to the last question should a priori be negative. In fact, the neurons possessing only one enzyme fail to synthesize MAs from the precursor amino acid. The existence of the MA transporter also cannot be considered as a specific marker of MA‐ergic neurons. In fact, some nonserotoninergic neurons possessing the serotonin transporter cannot synthesize serotonin from L‐tryptophan (De Vitry et al., 1986). The first attempt to improve the Swedish and Dutch classification of MA‐ergic systems has been made in eighties by mapping of the neurons expressing only AADC. The clusters of these neurons were called ‘‘D’’ groups (Jaeger et al., 1984). Although extensive data have been already accumulated in literature on the neurons partly expressing the MA‐ergic phenotype, their functional role, and the regulation remain far from complete understanding. According to the most promising hypothesis, the neurons partly expressing MA‐ergic phenotype serve to compensate a functional deficiency of MA‐ergic neurons (Ugrumov et al., 2002, 2004; Ershov et al., 2005) arising as a consequence of: (1) a transient decrease of the neuron functional activity; (2) a irreversible decrease of the neuron functional activity, e.g., under neurodegeneration; (3) an increased need of the brain in MAs that cannot be satisfied even by highly stimulated MA‐ergic neurons. The goal of this review is to evaluate the data concerning the location in the brain, development, regulation, and functional significance of the neurons partly expressing MA‐ergic phenotype in norm and pathology.

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Brain Neurons Partly Expressing the MA‐ergic Phenotype in Adult Mammals

2.1 Neurons Expressing Individual Enzymes of MA Synthesis Particular attention in literature and in this review is focused on the so‐called monoenzymatic neurons expressing individual enzymes of DA synthesis, TH or AADC that is explained by their particularly high frequency and wide distribution in the brain.

2.1.1 Hypothalamus The first attempt to estimate the frequency of monoenzymatic neurons has already shown that at least in the hypothalamus, one of the largest DA‐ergic center in the brain (A12, A13, A14), a number of TH‐ immunoreactive neurons (Van den Pol et al., 1984) greatly exceeded that of DA‐containing (fluorescent) neurons (Bjo¨rklund and Lindvall, 1984). It meant that numerous TH‐containing neurons lack AADC being incapable of DA synthesis. Later, the most interesting information about the monoenzymatic neurons was obtained when studying the hypothalamic arcuate nucleus (AN), suprachiasmatic nucleus (SCN), and the magnocellular supraoptic, paraventricular, and accessory nuclei. Arcuate nucleus. The initial immunocytochemical studies with mono‐immunostaining of the enzymes of DA synthesis have already shown incomplete overlapping in the distribution of TH‐immunoreactive neurons and AADC‐immunoreactive neurons in the AN in rats. The former were distributed all through the AN whereas the latter were mainly located dorsomedially (Okamura et al., 1988b). Similar distribution of TH‐immunoreactive neurons and AADC‐immunoreactive neurons is also a characteristic of the AN in other mammals including primates (Komori et al., 1991; Kitahama et al., 1998). It has been suggested that at least the ventrolateral portion of the AN contains monoenzymatic TH‐expressing neurons (Okamura et al., 1988b, c; Komori et al., 1991; Zoli et al., 1993) that was further proven by using double‐immunolabeling of the enzymes of DA synthesis (> Figures 2-3 and > 2-4) (Ershov et al., 2002a, b). Besides the ventrolateral region of the AN, the monoenzymatic TH‐neurons are located in

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. Figure 2‐3 Schematic drawing of the distribution of monoenzymatic TH‐neurons (filled circle), monoenzymatic AADC‐ neurons (opened circle), and bienzymatic (TH and AADC) neurons (asterisk) in the arcuate nucleus of adult rats. Arcuate nucleus is shown unilaterally. DM, dorsomedial region of the arcuate nucleus; VL, ventrolateral region of the arcuate nucleus

the rest of the nucleus. In contrast to monoenzymatic TH‐neurons, most monoenzymatic AADC‐neurons and bienzymatic neurons are located in the dorsomedial region of the AN, and only occasional neurons of either type are scattered in the ventrolateral region (> Figures 2‐3 and > 2‐4) (Ershov et al., 2002a). A population of monoenzymatic TH‐neurons is twice as small as the population of monoenzymatic AADC‐ neurons and the population of bienzymatic neurons in the AN of adult rats (Ershov et al., 2002a). From these quantitative data, it indirectly follows that relatively large populations of monoenzymatic neurons may be of functional importance. Suprachiasmatic nucleus. Besides AN, the SCN, a pacemaker of the circadian rhythmic activity (Klein et al., 1991) is a promising model for the study of monoenzymatic neurons. It has been demonstrated with double‐immunostaining in all studied mammals (rats, hamsters, sheep, shrews, and humans) that the SCN contains numerous monoenzymatic AADC‐neurons (D13) (Jaeger et al., 1983; Battaglia et al., 1995; Kitahama et al., 1998; Ishida et al., 2002) and occasional monoenzymatic TH‐neurons (Battaglia et al., 1995). The former population is located in the ventral and ventrolateral regions of the nucleus whereas the latter population is distributed at the periphery of the nucleus (> Figure 2-5) (Novak and Nunez, 1998). Hypothalamic ‘‘magnocellular’’ nuclei. Since the mid‐eighties, it has been repeatedly demonstrated that vasopressinergic neurons of the hypothalamic ‘‘magnocellular’’ nuclei, the supraoptic nucleus, the dorsolateral paraventricular nucleus, and ‘‘accessory’’ nuclei, are capable to coexpress TH in all studied mammals, rodents, and primates. In contrast to rodents, in humans TH is expressed not only in magnocellular vasopressinergic neurons but also in magnocellular oxytocinergic neurons (Panayotacopoulou et al., 1994). Rare magnocellular neurons coexpress TH in norm whereas their number multiplies under functional stimulation (> Figure 2-6) (Kiss and Mezey, 1986; Abramova et al., 2002). TH synthesized in neuronal cell

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. Figure 2‐4 The number of monoenzymatic TH‐expressing neurons (TH), monoenzymatic aromatic L‐amino acid decarboxylase (AADC)‐expressing neurons, and bienzymatic (TH and AADC) neurons in the ventrolateral (dotted line) and dorsomedial (solid line) regions of the arcuate nucleus (AN) of male rats at the 21st embryonic day (E21), the ninth postnatal day (P9) and in adulthood (Ershov et al., 2002a). Mean SEM. *P < 0.05

bodies is transported via axons and accumulated in axonal terminals in the pituitary posterior lobe (Abramova et al., 2000). Up to now, all the attempts to detect AADC in vasopressinergic neurons were unsuccessful (Kitahama et al., 1998). Other hypothalamic regions. In rodents, monoenzymatic TH‐neurons were detected in the periventricular nucleus, near the SCN (Battaglia et al., 1995), in the lateral preoptic area (Vincent and Hope, 1990), and the zona incerta (Skagerberg et al., 1988) whereas the monoenzymatic AADC‐neurons were found in the dorsomedial nucleus (D12) (Jaeger et al., 1984), premamillary nucleus (D8) (Jaeger et al., 1984; Karasawa et al., 1994; Kitahama et al., 1998), zona incerta (D10) (Skagerberg et al., 1988), lateral hypothalamus (D11), and in the medial preoptic area (Vincent and Hope, 1990; Kitahama et al., 1998).

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. Figure 2‐5 Schematic representation of the distribution of tyrosine hydroxylase‐immunopositive neurons (a) and aromatic L‐amino acid decarboxylase‐immunopositive neurons (b) in the hamster suprachiasmatic nuclei (SCN) (Novak and Nunez, 1998). The neurons are represented as dots. OC, optic chiasma; III, third ventricle

2.1.2 Striatum Rodents. Although dozens or even hundreds of immunocytochemical studies have been devoted to the nigrostriatal DA‐ergic system in intact mammals, only few of them described the neurons (cell bodies) containing enzymes of DA synthesis in the striatum and in the near limbic system, mostly nucleus accumbens in normal rodents and primates (Tashiro et al., 1989a, b; Betarbet et al., 1997; Mura et al., 2000; Lopez‐Real et al., 2003). In rats, the number of AADC‐immunoreactive neurons did not exceed 20 per striatum (Tashiro et al., 1989a) whereas TH‐immunoreactive neurons were either as numerous as AADC‐ neurons in number or even fewer (Tashiro et al., 1989b; Lopez‐Real et al., 2003). Besides the striatum, the TH‐immunoreactive neurons (1–3 per 40‐mm thick section) were found with mono‐immunolabeling in the nucleus accumbens of rodents. In most studies, no zonality in the distribution of the striatal monoenzymatic neurons has been recognized, though according to some authors the AADC‐immunoreactive neurons were mainly located in the subcallosal area of the dorsomedial, dorsal, and the dorsolateral striatum (Lopez‐ Real et al., 2003). Most TH‐immunoreactive neurons and AADC‐immunoreactive neurons had a round or oval cell body, which ranged from 10 to 20 mm in diameter, and extended 1–2 spiny processes, whereas rare neurons possessed several spiny dendrites (Tashiro et al., 1989a, b, 1990). Monkeys. In the primates (monkeys and humans), the number of the striatal neurons expressing enzymes of DA synthesis greatly exceeded that in the rodents (Betarbet et al., 1997; Cossette et al., 2005a). In normal

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. Figure 2‐6 Vasopressinergic neurons containing tyrosine hydroxylase, peptide (a, b) and mRNA (c, d), in supraoptic nucleus of intact (a, c) and salt‐loaded (b, d) adult rats (modified from Abramova et al., 2002). OT, optic tract; SON, supraoptic nucleus

monkeys (rhesus, macaques), TH‐immunoreactive neurons detected with mono‐immunolabeling were found in the caudate nucleus and putamen, mostly at their periphery (Dubach et al., 1987; Betarbet et al., 1997; Tande´ et al., 2006). Dorsally, the neurons were concentrated toward the dorsal border of the striatum near the corpus callosum. Ventrally, the striatal distribution of TH‐immunoreactive neurons appeared to be continuous with DA‐ergic neuron populations in the ventrobasal regions of the forebrain. Along the lateral edge of the putamen, the neurons were present in the neuropil and in the adjacent white matter. A few neurons were also located within the anterior limb of the internal capsule where it separated the caudate from the putamen. The AADC‐immunoreactive neurons (mono‐labeling) were distributed sparsely in the caudate nucleus, putamen, and the nucleus accumbens in the monkeys (Ikemoto et al., 1998a). Most of the TH‐immunoreactive neurons were round or oval in shape and small in size (10–18 mm) having from two to five primary aspiny processes. A few ( Figure 2-7) (Cossette et al., 2005a; Huot et al., 2007). Moreover, a small number of TH‐neurons were found in ventral margin of the nucleus accumbens (Ikemoto et al., 1998a). The most frequent TH‐immunoreactive neurons (58%) have a medium‐sized (diameter 20–24 mm) or small‐sized (10–15 mm) cell bodies with 3–4 varicose aspiny dendrites (Ikemoto et al., 1998a; Cossette et al., 2005a; Huot et al., 2007). Although aspiny TH‐neurons are distributed throughout the rostrocaudal extent of the caudate nucleus and the putamen, they predominate in the ventral striatum (> Figure 2-7). The

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. Figure 2‐7 Schematic representation of the distribution of tyrosine hydroxylase‐immunoreactive aspiny (circle) and spiny (star) neurons on frontal sections through the human basal ganglia in rostrocaudal (a, b) extension (Cossette et al., 2005a). AC, anterior commissure; CD, caudate nucleus; CL, claustrum; GPe, external segment of the globus pallidus; GPi, internal segment of the globus pallidus; IC, internal capsule; NA, nucleus accumbens; PUT, putamen; LV, lateral ventricle

minor population of TH‐IR neurons is represented by large neurons (6.5%) (25–35 mm) with spiny dendrites and few TH‐immunoreactive cells displaying mixed neuron–glial morphology. The latter have round or ovoid cell body (10–12 mm) with up to 10 aspiny processes. In contrast to aspiny TH‐IR neurons, the spiny TH neurons were detected in the caudate nucleus but not in the putamen (> Figure 2-7). The TH neurons with neuron–glial morphology are confined to the striatal bridges that link the caudate nucleus and the putamen rostrally. A single human striatum contains in average 331 TH‐immunoreactive neurons. However this number could have been increased by about 10 times if the prolific zone, containing about 3000 cells, observed in some brains had been taken into account (Cossette et al., 2005a). According to the mono‐immunolabeling study, rather numerous AADC‐immunoreactive neurons were distributed in the entire rostrocaudal extent of the striatum and the nucleus accumbens (> Figure 2-8) though their number varies significantly from human to human. The most numerous AADC‐neurons were located in the putamen (
120 neurons per 50 mm thick section), less numerous ones were situated in the nucleus accumbens (
90 neurons per 50 mm thick section), and the least number of AADC‐ immunoreactive neurons was observed in the caudate nucleus (
30 neurons per 50 mm thick section) (Ikemoto et al., 2003). Most AADC‐neurons are fusiform, bipolar, or multipolar in shape, medium to large in size (15–30 mm) having thick dendritic arbors (Ikemoto et al., 1997).

2.1.3 Other Brain Regions Cortex. Rather numerous monoenzymatic TH‐neurons and AADC‐neurons were detected with the double‐ immunolabeling of TH and AADC in the cortex, e.g., in the human anterior cingulate cortex (Ikemoto et al., 1999). These neurons lack VMAT2 (Weihe et al., 2006). The certain zonality in the distribution of TH‐neurons within the cortex has been recognized: the highest concentration is a characteristic of multi‐ associative cortex, a moderate concentration is typical for the unimodal associative cortex, and a minor concentration is an attribute of the visual, sensory, and motor primary areas (Gaspar et al., 1987). Substantia nigra and the ventral tegmental area. In addition to numerous DA‐ergic neurons innervating the striatum, the compact zone of the substantia nigra contains rare monoenzymatic TH‐neurons and AADC‐neurons that was shown initially in humans (Ikemoto et al., 1998b). In contrast to the substantia

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. Figure 2‐8 Schematic representation of the distribution of aromatic L‐amino acid decarboxylase‐immunoreactive neurons (dots) in the human striatum (Ikemoto et al., 1997). Acc, nucleus accumbens; Cn, caudate nucleus; Pu, putamen

nigra, rather numerous monoenzymatic TH‐neurons and AADC‐neurons were found in the ventral tegmental area (Ikemoto et al., 1998b). Other brain regions. Besides the hypothalamus, cortex, and the striatum, monoenzymatic TH‐neurons were found in the basal ganglia and in the globus pallidus in particular that was initially shown in the humans (Komori et al., 1991). Moreover, the clusters of monoenzymatic TH‐neurons were detected in hamsters in the nucleus of the horizontal limb of the diagonal band, pericentral nucleus of the inferior colliculus, lateral parabrachial nucleus, the dorsal motor nucleus of the vagus, and the substantia innominata, beneath the rostral globus pallidus (Vincent and Hope, 1990). Rare TH‐neurons are located in the anterior olfactory nucleus in mice (Nagatsu et al., 1990) and rats (Meredith et al., 1999). The monoenzymatic AADC‐neurons were observed in rodents in the anterior olfactory nucleus, pretectal nucleus (D5), the nucleus of the solitary tract (D2) (Karasawa et al., 1994), and in the internal division of the lateral parabrachial nucleus (Vincent and Hope, 1990). Moreover, a small number of TH‐ immunoreactive and AADC‐immunoreactive Purkinje cells were detected with mono‐immunostaining in the cerebellum (Sakai et al., 1995). Taking into account that the number of TH‐immunoreactive neurons is twice as large as AADC‐immunoreactive neurons, at least half of TH‐immunoreactive neurons should be monoenzymatic in nature (Sakai et al., 1995).

2.2 Bienzymatic TH- and AADC-Expressing Neurons According to the double‐immunolabeling studies, the striatum of rodents, monkeys, and humans contains bienzymatic (TH and AADC) neurons in addition to the monoenzymatic ones (Ikemoto et al., 1998a; Weihe et al., 2006). However, in contrast to true DA‐ergic neurons, they lack VMAT2 (Weihe et al., 2006). The bienzymatic neurons are distributed along the ventral margin of the rostral nucleus accumbens and the caudate nucleus. They were small‐ to medium‐sized (10–15 mm), elongated in shape, having one or two processes (Ikemoto et al., 1998a).

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2.3 Non-MA-ergic Neurons Expressing the MA Transporters Up to the present, no special studies of the expression of the MA transporters in non‐MA‐ergic neurons have been made by using multi‐labeling of at least two first enzymes of MA (DA or serotonin) synthesis and the MA transporter. Nevertheless, according to indirect evidence, at least two types of non‐MA‐ergic neurons expressing the MA transporter may be recognized: the monoenzymatic neurons and the neurons lacking enzymes of MA synthesis. Some neurons express the serotonin transporter while others – the DA transporter (> Figures 2-9 and > 2-10) (Lorang et al., 1994; Hoffman et al., 1998).

. Figure 2‐9 Neurons containing either tyrosine hydroxylase or dopamine transporter mRNA on successive coronal levels of the rat hypothalamus in rostrocaudal extension (a–d) (Lorang et al., 1994). Open squares, TH‐immunoreactive neurons; open circles, DAT mRNA‐containing and TH‐immunonegative neurons; solid circles, double‐labeled (TH‐immunoreactive, DAT mRNA‐containing) neurons. AH, anterior hypothalamic nucleus; AN, arcuate nucleus; DM, dorsomedial nucleus; LPA, lateral preoptic area; ME, median eminence; MPA, medial preoptic area; OC, optic chiasm; PV, paraventricular nucleus; VM, ventromedial nucleus; III, third ventricle; ZI, zona incerta

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. Figure 2‐10 Schematic representation of the distribution of neurons containing vesicular monoamine transporter 2 mRNA (left hemisphere of each level) or serotonin (open square) and dopamine (filled triangles) transporter mRNAs (right hemisphere of each level) at coronal successive levels in rostrocaudal extension of the adult rat hypothalamus (Hoffman et al., 1998). Open circles, overlap of vesicular monoamine transporter 2 mRNA with either serotonin transporter or dopamine transporter; filled circles, neurons expressing vesicular monoamine transporter 2 mRNA which do not correspond to neurons expressing either serotonin transporter or dopamine transporter mRNAs. AC, anterior commissure; AH, anterior hypothalamic nucleus; AN, arcuate nucleus; DM, dorsomedial nucleus; DP, dorsal premamillary nucleus; LH, lateral hypothalamus; PH, posterior hypothalamic nucleus; MP, medial preoptic area; MV, mamillary nucleus, ventral part; OC, optic chiasm; PV, paraventricular nucleus; VM, ventromedial nucleus; VP, ventral premamillary nucleus; SC, suprachiasmatic nucleus; ZI, zona incerta; III, third ventricle

Neurons expressing the serotonin transporter. Up to the present, the only one population of non‐MA‐ ergic neurons expressing a serotonin transporter has been recognized with certainty. These small oval mono‐ and bipolar neurons with short unbranched processes were detected in the hypothalamic dorsomedial nucleus in rats. They contain serotonin transporter mRNA (> Figure 2-10) (Hoffman et al., 1998) and the functionally active protein that is manifested by: (1) the radiolabeling of the neurons following 3 H‐serotonin injections to the cerebral ventricles (Beaudet and Descarries, 1979; Ugrumov et al., 1986);

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(2) the appearance of the serotonin fluorescent neurons after the intraventricular injections of serotonin (Fuxe and Ungerstedt, 1968); and (3) the neuron degeneration provoked by 5,7‐dihydroxytryptamine, the neurotoxin of serotoninergic neurons (Frankfurt and Azmitia, 1983). These neurons are not serotoninergic in nature since they lack tryptophan hydroxylase, the first rate‐limiting enzyme of serotonin synthesis, and the presence of AADC is questionable (Frankfurt et al., 1981; Ugrumov et al., 1989a). Similar neurons were found in the substantia nigra, the ventral tegmental area, the nucleus sagulum, the Barrington nucleus, and the parvicellular reticular nucleus (Hoffman et al., 1998). Neurons expressing the DA transporter. Although there is no convincing evidence of the existence of non‐ MA‐ergic neurons expressing the DA transporter, some data may be interpreted in favor of this idea. One can state that at least a part of the striatal monoenzymatic TH‐neurons coexpress the DA transporter since it is colocalized in the whole population of striatal TH‐immunoreactive neurons (Porritt et al., 2000; Cossette et al., 2005b; Tande´ et al., 2006) composed of monoenzymatic TH‐neurons and bienzymatic (TH and AADC) neurons (Lopez‐Real et al., 2003). In this context, the author’s statement that all the TH‐immunoreactive neurons coexpressing the DA transporter are obligatory DA‐ergic in nature (Porritt et al., 2000; Cossette et al., 2005a, b; Tande´ et al., 2006) appears to be doubtful. In addition to monoenzymatic TH‐neurons, the DA transporter is coexpressed in the neurons lacking TH. Since the authors did not use a triple‐labeling technique, the DA transporter coexpression in monoenzymatic AADC‐neurons cannot be excluded. The neurons possessing the DA transporter but lacking TH have been detected in the zona incerta (> Figure 2-9), ventral premamillary nucleus (Lorang et al., 1994), Barrington nucleus, and in the pontine tegmentum (Hoffman et al., 1998).

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Brain Neurons Partly Expressing MA‐ergic Phenotype in Mammals in Ontogenesis

Major data on the monoenzymatic neurons in the brain in ontogenesis are rather fragmentary with an exception of the developing hypothalamus and the AN in particular.

3.1 Neurons Expressing Individual Enzymes of MA Synthesis 3.1.1 Hypothalamus Arcuate nucleus. According to the mono‐immunolabeling data, TH‐immunoreactive neurons first appear in the developing AN in rats on the 17–18th day of prenatal life (Daikoku et al., 1986; Ugrumov et al., 1989b). In the initial studies, these neurons were mistakenly considered as DA‐ergic neurons based on the conventional Swedish classification (Daikoku et al., 1986; Ugrumov et al., 1989b). Later, it has been shown by using mono‐immunostaining of TH and AADC on adjacent mirror sections that only TH is expressed in the neurons of the AN on the 18th fetal day (Balan et al., 2000). At that time and later, on the 20–21st embryonic day, most monoenzymatic TH‐neurons were located in the ventrolateral region of the nucleus (> Figure 2-11) (Balan et al., 2000). In contrast to TH‐immunoreactive neurons, the AADC‐immunoreactive neurons first appear in the rat AN on the 20th fetal day. Since this time onward, monoenzymatic AADC‐immunoreactive neurons are located mainly in the dorsomedial region of the nucleus (> Figure 2-11) being in perinatal rats as numerous as monoenzymatic TH‐immunoreactive neurons (> Figure 2-4). Subsequent application of the double‐ immunolabeling of TH and AADC has shown that the AN of rat fetuses contains more than 99% monoenzymatic TH‐neurons and AADC‐neurons (1:1), and less than 1% bienzymatic neurons at the end of prenatal life (> Figure 2-12) (Balan et al., 2000; Ershov et al., 2002a). The total number of monoenzymatic TH‐neurons, monoenzymatic AADC‐neurons, and bienzymatic DA‐ergic neurons in the AN was about 850, 850, and 17, respectively (> Figure 2-4). From the ninth postnatal day to adulthood, the number of monoenzymatic TH‐neurons decreases significantly (
430 neurons per nucleus), the number of monoenzymatic AADC‐neurons remains at the

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. Figure 2‐11 Tyrosine hydroxylase‐immunoreactive neurons (a) and aromatic L‐amino acid decarboxylase‐immunoreactive neurons (b) (Balan et al., 2000); schematic representation of their distribution in the arcuate nucleus of rat fetuses on the 21st day of intrauterine development. AN, arcuate nucleus; DM, dorsomedial region of the AN; ME, median eminence; VL, ventrolateral region of the AN; III, third ventricle

same level as in fetuses (
850 neurons per nucleus), and the number of bienzymatic neurons greatly increases (
1200 neurons per nucleus). However in adulthood, the monoenzymatic neurons are still as numerous as the bienzymatic ones (> Figure 2-4) (Ershov et al., 2002a). Although the monoenzymatic and bienzymatic neurons are distributed all through the AN in postnatal rats, most monoenzymatic TH‐neurons are located in the ventrolateral region whereas the monoenzymatic AADC‐neurons and bienzymatic neurons are concentrated in the dorsomedial region (> Figures 2-4 and > 2-12) (Ershov et al., 2002a). From the end of prenatal life to adulthood, the monoenzymatic TH‐neurons, AADC‐neurons and bienzymatic neurons were often seen in close topographic relations (Ershov et al., 2002a). According to confocal microscopy, the neurons containing enzymes of DA synthesis are in appositions both at the level of the distal axons in the median eminence and at the level of cell bodies in the AN (> Figures 2-13 and > 2-14) (Ershov et al., 2002b). The frequency of these contacts increases gradually with age, mainly due to the overgrowth and ramification of the neuron processes. In the previous ontogenetic studies, the fibers mono‐immunostained for TH or possessing histofluorescent DA detected in the developing median eminence have a priori been considered as bienzymatic DA‐ergic in nature (Ibata et al., 1982; Ugrumov et al., 1989c). A subsequent application of the double‐ immunostaining of the enzymes of DA synthesis first made it possible to change this consideration by evaluating the ingrowth of monoenzymatic and bienzymatic fibers to the median eminence in ontogenesis, separately. The density of monoenzymatic TH‐axons, i.e., the number of axons per unit of the volume of the

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. Figure 2‐12 The number of monoenzymatic tyrosine hydroxylase‐immunoreactive neurons (TH), monoenzymatic aromatic L‐amino acid hydroxylase‐immunoreactive neurons (AADC), and bienzymatic tyrosine hydroxylase‐ immunoreactive and aromatic L‐amino acid hydroxylase‐immunoreactive neurons (TH and AADC) in the arcuate nucleus of rat fetuses on the 21st day of intrauterine development (Ershov et al., 2002a). *P < 0.05

median eminence is maximal at the end of prenatal life whereas it decreases gradually during postnatal period (> Figure 2-15) (Ershov et al., 2002b). The age variation in the density of monoenzymatic TH‐axons might be a consequence of: (1) coexpression of AADC in the neurons initially synthesizing only TH; (2) establishment of either neural or hormonal inhibitory controls of TH synthesis (see > Section 8); (3) apoptosis of some monoenzymatic TH‐neurons. However, it should be emphasized that the decrease of the density of monoenzymatic TH‐axons in the median eminence with age does not obligatory mean a decrease of their total number as the volume of the median eminence enlarges significantly in rats during postnatal period (Ugrumov et al., 1985a). In contrast to monoenzymatic TH‐axons, the density of monoenzymatic AADC‐axons increases progressively from the end of fetal life to adulthood (> Figure 2-15) (Ershov et al., 2002b) though the number of monoenzymatic AADC‐neurons does not change over the same period (> Figure 2-4) (Ershov et al., 2002a, b). This seeming controversy might be explained by: (1) an intense ramification of the axonal projections of monoenzymatic AADC‐neurons of the AN; (2) the ingrowth of monoenzymatic AADC‐ axons to the median eminence from the outside of the AN (Jaeger et al., 1984). The density of bienzymatic axons increases gradually during pre‐ and postnatal period that correlates well with the increasing number of bienzymatic neurons in the AN (> Figures 2-4 and > 2-15). From this correlation it indirectly follows that a great portion of bienzymatic axons in the median eminence belongs to the neurons of the AN. Suprachiasmatic nucleus. Although the information about the monoenzymatic neurons in the developing SCN is quite limited, it is known that it contains monoenzymatic AADC‐neurons in young rats (P10), the only studied period of ontogenesis, as in adulthood (Battaglia et al., 1995). Moreover, rare TH‐immunoreactive most probably monoenzymatic neurons were first detected with the electron microscopic immunocytochemistry in the rat SCN even earlier, at the end of prenatal life. From this time onward, they are located in the periphery of the nucleus (Ugrumov et al., 1994).

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. Figure 2‐13 Schematic representation of topographic relations between monoenzymatic tyrosine hydroxylase (TH)‐neurons and monoenzymatic aromatic L‐amino acid decarboxylase (AADC)‐neurons in the arcuate nucleus and median eminence as well as of hypothetical L‐DOPA transfer from the former to the latter (Ugrumov et al., 2002). BV, blood vessels of the hypophysial portal circulation; DA, dopamine

The data on the transient innervation of the SCN with the monoenzymatic TH‐fibers in ontogenesis appears to be of particular interest (Ugrumov et al., 1989c; Beltramo et al., 1994). Indeed, it has been demonstrated that the ventral and ventromedial portions of the SCN are intensely innervated by TH‐immunoreactive fibers in young rodents (> Figure 2-16) (Ugrumov et al., 1989c) that is in contrast to adults. Irrespectively of the sex, TH‐immunoreactive fibers first appear in the rat SCN on the second postnatal day. Thereafter, the number of fibers increases rapidly until reaching maximum at the 10th postnatal day. Then, the fiber density decreases rapidly (> Figure 2-16) up to puberty (Beltramo et al., 1994). TH‐immunoreactive fibers occurred to be non‐MA‐ergic in nature as they lack AADC (Battaglia et al., 1995) and transporters of catecholamines and serotonin. Indeed, 6‐hydroxydopamine (6‐OHDA) and 5,7‐ dihydroxytryptamine, neurotoxins of DA‐ergic and serotoninergic neurons, which are captured by the MA transporters, did not provoke the degeneration of TH‐immunoreactive fibers (> Figure 2-16) (Beltramo et al., 1994). Most TH‐immunoreactive fibers innervating the SCN belong to a loose accumulation of small monoenzymatic TH‐neurons scattered in the vicinity of the SCN, mainly in the ventral portion of the anterior hypothalamic nucleus (Mirochnik et al., 2002). These TH‐immunoreactive neurons first appear in rats just after birth, followed by their continuous increase in number for subsequent two weeks. Then, the population of TH‐immunoreactive neurons diminishes rapidly and disappears by puberty, synchronously with the disappearance of the monoenzymatic TH‐fibers in the SCN. The axons of TH‐immunoreactive neurons of

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. Figure 2‐14 Axo‐somatic specialized‐like contact (arrowhead) between monoenzymatic tyrosine hydroxylase (TH)‐immunoreactive axon (light grey) and monoenzymatic aromatic L‐amino acid decarboxylase (AADC)‐immunoreactive cell body in the arcuate nucleus of the prepubertal rat (Ershov et al., 2002b). Confocal microscopy: double‐labeling of TH and AADC (a) and mono‐labeling of AADC (b) and TH (c). Dotted line, outlined AADC‐immunoreactive neuron

this location are often oriented toward the SCN, and one may follow some of them on thick sections up to the SCN. In addition to monoenzymatic TH‐neurons in the anterior hypothalamic nucleus, rare TH‐ neurons located in the periphery of the SCN or in the next periventricular nucleus (Battaglia et al., 1995) appear to contribute to its transient innervation. Hypothalamic magnocellular nuclei. In ontogenesis, TH is initially expressed in the magnocellular vasopressinergic neurons of rats at the end of the second postnatal week. This is manifested by the appearance of rare neurons possessing TH‐immunoreactive material and TH mRNA in the supraoptic nucleus (Ugrumov, 2002). In contrast to rodents, in humans, TH is expressed not only in differentiating vasopressinergic neurons but also in oxytocinergic neurons (Panayotacopoulou et al., 1994).

3.1.2 Extrahypothalamic Regions The neurons which are TH‐immunoreactive transiently over certain periods of ontogenesis were recognized in many areas of the developing brain. First TH‐immunoreactive neurons increase in number reaching the peak, and then they decrease gradually in number and finally disappear. The transient neuron populations of this kind were found: (1) in the raphe nucleus of the laboratory shrew over the first two weeks of postnatal life (Karasawa et al., 1997); (2) in the anterior olfactory nucleus of mice from the 16th fetal day to puberty with the plateau during the second postnatal week (Nagatsu et al., 1990); (3) in the olfactory bulbs in fetuses of rats and humans (Verney et al., 1996; Izvolskaia et al., 2006); (4) in the limbic system, the bed

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. Figure 2‐15 Density of monoenzymatic TH‐immunoreactive fibers (TH), monoenzymatic aromatic L‐amino acid decarboxylase‐ immunoreactive fibers (AADC), and bienzymatic (TH&AADC) fibers in the median eminence of rats at the 21st embryonic day (E21), the ninth postnatal day (P9) and in adulthood (Ershov et al., 2002b). Mean SEM. *P < 0.05

nucleus of the stria terminalis, and the central nucleus of amygdala in rats from the 17th prenatal day to puberty with the maximal concentration in the early postnatal period (Verney et al., 1988); (5) in the medial geniculate nucleus in mice from the first to the fourth postnatal week with the maximal number during the third week of life (Nagatsu et al., 1996); (6) in the neocortex in rats for the first three weeks of postnatal life (Berger et al., 1985), in the striatum in rat fetuses (> Figure 2-17) (Sorokin et al., unpublished). Some differentiating TH‐neurons synthesize classical neurotransmitters, serotonin (Karasawa et al., 1997), or neuropeptides: gonadotropin‐releasing hormone, somatostatin, and substance P (Verney et al., 1988, 1996; Izvolskaia et al., 2006). According to the double‐immunolabeling studies, differentiating TH neurons do not coexpress AADC (Berger et al., 1985; Verney et al., 1988; Nagatsu et al., 1990, 1996), apparently except the serotoninergic neurons of the raphe nucleus (Karasawa et al., 1997). According to the author’s opinion, the disappearance of TH‐immunoreactive neurons in ontogenesis is explained by: (1) the degeneration of the neurons coexpressing TH; (2) the turning off TH‐expression with age; and (3) the decrease of TH synthesis and hence of the intracellular content of the TH‐immunoreactive material under the level undetectable with immunocytochemistry (Karasawa et al., 1997). The studies of the monoenzymatic AADC‐neurons in ontogenesis are less numerous than those of monoenzymatic TH‐neurons. Nevertheless, it has been shown that between fourteen D groups recognized in adults, nine groups (D1, D4–D7, D10–D12, and D14) first appear in rats during prenatal period, whereas the remaining five groups become detectable with immunocytochemistry during the first postnatal week (Jaeger and Teitelman, 1992). There is a ventrodorsal gradient in the appearance of AADC‐immunoreactive neurons in the developing brain: the neurons of the ventral groups attain AADC‐immunoreactivity prior to the neurons of the dorsal groups.

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. Figure 2‐16 Strong innervation of the suprachiasmatic nucleus (SCN) in rats at the tenth postnatal day by tyrosine hydroxylase(TH)‐immunoreactive fibers (a, b) (Ugrumov et al., 1989c) which are resistant to 6‐hydroxydopamine (6‐OHDA) (C, left side) that is in contrast to tyrosine hydroxylase‐immunoreactive fibers of the paraventricular nucleus (PVN) (C, right side) in the same animals (c) (Beltramo et al., 1994). *P < 0.05

. Figure 2‐17 Tyrosine hydroxylase-immunoreactive neurons (arrows) in the striatum and neural epithelium around the lateral ventricles of rat fetuses on the 21st day of intrauterine development (Sorokin et al., unpublished)

Monoenzymatic AADC‐immunoreactive neurons first appear in the rat brain at the 15th embryonic day. At that time, all the neurons are gathered together representing a large cluster located in the intermediate region of the lateral diencephalic vesicle. Later, this accumulation gives rise first to ventral groups located in the lateral hypothalamic area (D11) and in the anterior hypothalamus, medial to the medial forebrain bundle (D14), and then to dorsal groups distributed in the dorsal mesencephalon (D5) and in the mid‐hypothalamus in its rostrocaudal extent. Rostrally, this group extends to the level of the bed nucleus segregating into the most anterior cluster adjacent to the lateral preoptic area (D11) and the accumulation located just beyond the anterior commissure (D14). From the 16th to the 19th prenatal day, a number of additional clusters of monoenzymatic AADC‐immunoreactive neurons become visible. They are

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located in the dorsal mesencephalon (D4) and the dorsal diencephalon (D6, D7, and D10), as well as in the dorsomedial nucleus (D12), thalamus (D7), and the lateral habenular nucleus (D6). From the earlier data it follows that most accumulations of AADC‐neurons appear in rats by the end of intrauterine development, except groups D2, D3, D8, D9, and D13 which appear during the early postnatal period (Jaeger and Teitelman, 1992).

3.2 Non-MA-ergic Neurons Expressing the MA Transporters Neurons partly expressing the serotoninergic phenotype, the high‐affinity serotonin uptake, but lacking either both enzymes of serotonin synthesis or tryptophan hydroxylase have been discovered in the rat diencephalon in ontogenesis (Ugrumov et al., 1986, 1989a) as in adulthood (see, > Section 3.3). In contrast to adult rats, in fetal and young rats these neurons are more widely distributed in the forebrain, and transiently express the serotonin transporter (Ugrumov et al., 1986, 1989a; Gaspar et al., 2003). The neurons of this kind were found in the hypothalamus: the dorsomedial nucleus, the SCN, and the preoptic area (> Figure 2-18), as well as in the thalamus, limbic cortex, retina, and superior olivary nucleus of rodents from the 15th embryonic day to the tenth postnatal day (Ugrumov et al., 1986, 1989a; Lebrand et al., 1998; Gaspar et al., 2003). The neurons are quite different in morphology even within the same brain area. Indeed, the neurons of the hypothalamic dorsomedial nucleus differed considerably in their appearance from those in the anterior

. Figure 2‐18 Schematic representation of the distribution of nonmonoaminergic neurons expressing serotonin transporter in the diencephalon of perinatal rats (Ugrumov et al., 1986, 1989c). AC, anterior commissure; DM, dorsomedial nucleus; LV, lateral ventricle; MFB, medial forebrain bundle; OC, optic chiasm; ON, optic nerve; PA, preoptic area; SC, suprachiasmatic nucleus; III, third ventricle

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hypothalamus. The former population was represented by uni‐ or bipolar neurons small in size (5–8 mm) and round in shape with slightly ramified processes, whereas the latter population composed of bipolar neurons larger in size (6  8  15 25 mm) and oval in shape with long widely ramified processes (> Figure 2-19) (Ugrumov et al., 1986, 1989a). . Figure 2‐19 Neurons expressing serotonin transporter (arrows) in the preoptic area of rat fetuses (18th fetal day) (a, b) and in dorsomedial nucleus in young rats (tenth postnatal day) (c, d) at low (a, c) and high (b, d, e) magnifications. Neurons become serotonin‐immunoreactive after systemic treatment with serotonin precursors (a–d) and capture specifically intraventricularly injected tritiated serotonin (E). Both types of labeling were prevented by preliminary injection of inhibitor of serotonin uptake (Ugrumov et al., 1986, 1989a). DM, dorsomedial nucleus; PA, preoptic area; III, third ventricle

The neurons of the SCN and the dorsomedial nucleus possessing the serotonin transporter have been detected in fetal and neonatal rats with autoradiography following the 3H‐serotonin intraventricular injection (Ugrumov et al., 1986), whereas the existence of non‐MA‐ergic neurons expressing the serotonin transporter in the lateral preoptic area was proven by using a combination of immunocytochemistry for serotonin and a pharmacological approach. The latter became detectable with immunocytochemistry for serotonin but only after the animal treatment with L‐tryptophan or 5‐hydroxytryptophan, serotonin precursors, and pargyline, the inhibitor of MA oxidase. However, this effect has been abolished by the preliminary treatment with fluoxetine, an inhibitor of the serotonin uptake (Ugrumov et al., 1989a). According to the author’s interpretation, the pretreatment with the serotonin precursor stimulated serotonin synthesis and release in serotoninergic neurons that was followed by the serotonin uptake from the extracellular space to nonserotoninergic neurons expressing the serotonin transporter (Ugrumov et al., 1989a). In addition to the in vivo studies, the hypothalamic neurons possessing the serotonin transporter have been detected in the primary tissue culture of the embryonic hypothalamus (De Vitry et al., 1986). Although a number of studies have provided indirect evidence of the existence of the non‐MA‐ergic neurons expressing the DA transporter in the brain of adult mammals, the information about the neurons of this kind in ontogenesis is not still available.

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Functional Properties and Functional Significance of the Neurons Partly Expressing the MA‐ergic Phenotype

The discovery of non‐MA‐ergic neurons expressing individual enzymes of MA synthesis has raised a question about their functional properties and functional significance.

4.1 Monoenzymatic Neurons Expressing TH TH is represented in monoenzymatic neurons by the same isoforms (Marsais et al., 2002) as in DA‐ergic neurons of the substantia nigra (Nagatsu, 1995). Furthermore, in most monoenzymatic neurons, TH is enzymatically active providing L‐DOPA synthesis from L‐tyrosine as a final synthetic product. This was proven by a number of observations made in the brain areas possessing mostly monoenzymatic TH‐ neurons: (1) the neurons being immunoreactive for L‐DOPA but not for DA (Meister et al., 1988; Okamura et al., 1988a, b, c); (2) the neurons lacking DA histofluorescence even after systemic administration of exogenous L‐DOPA (Zoli et al., 1993); and (3) the neurons showing a high level of L‐DOPA synthesis (> Figure 2-20) (Melnikova et al., 1999; Ugrumov et al., 2002). The neurons with L‐DOPA synthesis as a final synthetic product were detected with certainty in the AN, substantia nigra, ventral tegmental region, and the raphe nucleus (Mons et al., 1989). . Figure 2‐20 þ L‐DOPA concentration and K ‐stimulated release in the mediobasal hypothalamus (arcuate nucleus and median eminence) of rat fetuses at the 21st day of intrauterine development (Melnikova et al., 1999). Mean SEM. *P < 0.05

Nevertheless in some monoenzymatic neurons, TH fails to convert L‐tyrosine to L‐DOPA most probably because of the absence of tetrahydrobiopterin, the specific cofactor, synthesized with guanosine triphosphate (GTP) cyclohydroxylase I (Nagatsu et al., 1997). Indeed, the neurons of this kind: hypothalamic magnocellular vasopressinergic neurons (Marsais et al., 2002), cerebellar Purkinje cells (Sakai et al., 1995),

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the neurons of the developing anterior olfactory nucleus (Nagatsu et al., 1990), and the neurons of the geniculate nucleus (Nagatsu et al., 1996) are immunoreactive for TH, but immunonegative for L‐DOPA and GTP cyclohydroxylase I. Therefore, the Weihe et al. (2006) suggestion to call all the monoenzymatic TH‐neurons ‘‘dopaergic’’ neurons is incorrect. At least a part of monoenzymatic TH‐neurons does not coexpress the DA transporter. For instance, the monoenzymatic TH‐neurons in the ventrolateral portion of the AN lack the DA transporter mRNA and protein (> Figure 2-21) (Hoffman et al., 1998) that is in line with the biochemical observation of the low‐ affinity DA uptake in the whole AN (Moore et al., 1985; Melnikova et al., 1999). Similar conclusion has been

. Figure 2‐21 Tyrosine hydroxylase (TH), dopamine transporter (DAT) and vesicular monoamine transporter 2 (VMAT2) expression in the arcuate nucleus: TH expression in both dorsomedial (arrow) and ventrolateral (arrowhead) regions of the arcuate nucleus (a); DAT expression in the dorsomedial but not in the ventrolateral region (b); VMAT2 expression in the dorsomedial region (c) (Hoffman et al., 1998). ME, median eminence; III, third ventricle

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made when studying the SCN of young rats (Beltramo et al., 1994). It has been shown that the monoenzymatic TH‐fibers innervated the SCN during the early postnatal period (Ugrumov et al., 1989c; Beltramo et al., 1994) lack the DA and serotonin transporters. This was proven by the failure of 6‐OHDA and 5,7‐dihydroxytryptamine, neurotoxins of DA‐ergic, and serotoninergic neurons, to provoke the degeneration of TH‐immunoreactive fibers (> Figure 2-16) (Beltramo et al., 1994). The TH‐immunoreactive neurons lacking the DA transporter are widely distributed in the brain, particularly in the diencephalon. They were found in the periventricular nucleus (A14), preoptic area (A15), supraoptic nucleus, and the posterior hypothalamus (A11), as well as outside the diencephalon in the bed nucleus of the stria terminalis (A15), rostral liner nucleus, rostral periaqueductal gray (A11), over the interpeduncular nucleus (> Figure 2-9) (Lorang et al., 1994). The authors probably made a mistake considering these TH‐immunoreactive neurons as DA‐ergic neurons (Lorang et al., 1994). That is why they found it difficult to interpret their functional significance being obliged to declare that DA produced by these ‘‘DA‐ergic’’ neurons provide a paracrine action on the target brain cells or an endocrine action on the pituitary lactrotrophes. This is probably not the true as the pituitaries of mice lacking the DA transporter fail to develop lactrotrophes (Bosse et al., 1997). The difficulty in interpretation of the functioning of diencephalic TH‐immunoreactive neurons lacking the DA transporter may be overcome if they are considered as monoenzymatic neurons producing L‐DOPA as a final synthetic product. Although L‐DOPA synthesis in most monoenzymatic TH‐neurons has been proven, the mechanism of its storage and release remains uncertain. The fact that all the monoenzymatic TH‐neurons irrespective of their location in the brain of the studied animals (rodents and monkeys) lack VMAT2 (> Figure 2-21) (Hoffman et al., 1998; Weihe et al., 2006), makes it questionable the intragranular accumulation of L‐DOPA. Nevertheless, it has been demonstrated ex vivo and in the primary cell culture of the AN of rat fetuses containing numerous monoenzymatic TH‐neurons but almost lacking bienzymatic DA‐ergic neurons (Ershov et al., 2002a) that a large amount of L‐DOPA is discharged under membrane depolarization (> Figure 2-20) (Melnikova et al., 1999). These data have definitely shown that the monoenzymatic TH‐neurons synthesize and store L‐DOPA which releases in response to the adequate physiological stimuli. L‐DOPA synthesis in the monoenzymatic TH‐neurons raised a question about its functional significance. Three possibilities are discussed in literature. According to the first suggestion, L‐DOPA plays a role of the intracellular signal controlling neuronal metabolism, including synthesis and release of non‐MA neurotransmitters and neuromodulators, mainly of neuropeptides. In fact, TH is coexpressed in peptidergic neurons: (1) permanently in adulthood, e.g., in the neurons of the AN producing GABA (g‐aminobutyric acid), neurotensin, somatostatin, growth hormone‐releasing hormone, dinorphin, and galanin (Everitt et al., 1986; Chaillou et al., 1998); (2) in adulthood transiently under certain physiological conditions, e.g., in magnocellular vasopressinergic neurons under functional stimulation (Yagita et al., 1994); (3) transiently during certain periods of ontogenesis that was shown for the differentiating neurons producing somatostatin and substance P (Verney et al., 1988). It should be mentioned that some authors have considered a priori peptidergic neurons coexpressing TH as catecholaminergic in nature though they did not attempt to detect AADC (Everitt et al., 1986). In addition to vasopressinergic neurons, TH but not AADC is probably coexpressed in a wide range of peptidergic neurons. This suggestion is strongly supported by the overlapping in the distribution of neurotensin‐, galanin‐, growth hormone‐releasing hormone‐, and corticotrophin‐producing neurons coexpressing TH with the monoenzymatic TH‐neurons in the ventrolateral part of the AN (Okamura et al., 1985; Everitt et al., 1986), which almost lacks monoenzymatic AADC‐neurons and bienzymatic DA‐ergic neurons (Ershov et al., 2002a). TH is colocalized in differentiating neurons not only with neuropeptides but also with classical neurotransmitters, e.g., serotonin, or the key enzymes of their synthesis, e.g., choline acetyltransferase (Tinner et al., 1989; Karasawa et al., 1997). The authors believe that these neurons produce L‐DOPA which contributes to the regulation of the neuron differentiation (Verney et al., 1996; Izvolskaia et al., 2006). However, all the attempts to detect L‐DOPA in differentiating noncatecholaminergic TH‐expressing neurons were unsuccessful (Karasawa et al., 1997). According to the second suggestion, L‐DOPA synthesized in monoenzymatic TH‐neurons plays a role of an intercellular signal like classical neurotransmitters or neuromodulators (Misu et al., 2003). The authors have raised a number of arguments in favor of this hypothesis: (1) Caþ2‐dependent mechanism of L‐DOPA

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release from the striatum that has been shown in vivo (Nakamura et al., 1992) and in vitro (Goshima et al., 1988); (2) stimulating effect of L‐DOPA via most probably presynaptic b‐adrenoreceptors on the noradrenaline release from the hypothalamic slices that has been shown under the pharmacological inhibition of AADC (Goshima et al., 1991); (3) dose‐dependent stimulating or inhibitory effects of L‐DOPA on the DA release from the DA‐ergic nigrostriatal system mediated most probably via presynaptic D2 receptors (Fisher et al., 2000); and (4) dose‐dependent inhibitory action of L‐DOPA on AADC activity and DA release in the DA‐ergic nigrostriatal system (Fisher et al., 2000). The observation of close topographic relations of monoenzymatic TH‐neurons with the presumptive targets for L‐DOPA, e.g., with the vasoactive intestinal polypeptide‐producing neurons in the SCN in young rats (Battaglia et al., 1995) may also be interpreted in favor of the earlier hypothesis. Although the hypothesis about the L‐DOPA functioning as a neurotransmitter is attractive, some arguments should be considered with caution. Indeed, some principal arguments supporting this concept have derived from the experiments with the AADC inhibition and the presumptive shutdown of catecholamine synthesis (Misu et al., 2003). In practice, it is quiet difficult to inhibit completely the enzymes of synthesis of classical neurotransmitters when using a pharmacological approach. The observation of monoenzymatic TH‐immunoreactive fibers (Ershov et al., 2002b) or the L‐DOPA‐ immunoreactive fibers abutting on the primary capillary plexus of the hypophysial portal circulation in the median eminence (Misu et al., 2003) is considered a morphological evidence of the L‐DOPA delivery from the monoenzymatic TH‐axons to the hypophysial portal circulation. In this particular case, L‐DOPA may play a role of a neurohormone reaching the pituitary via bloodstream and acting on the glandular cells as a neurohormone.

4.2 Monoenzymatic Neurons Expressing AADC In most studied monoenzymatic AADC‐neurons, AADC was shown to be capable of converting L‐DOPA to DA and 5‐hydroxytryptophan to serotonin (Karasawa et al., 1994; Ishida et al., 2002). Either precursor is known to be captured to monoenzymatic AADC‐neurons by the membrane transporter of large neutral amino acids (Sugaya et al., 2001; Ferna´ndez et al., 2005). DA synthesis in monoenzymatic AADC‐neurons has been proven by the appearance of DA‐histofluorescence or DA‐immunoreactive materials after the systemic administration of exogenous L‐DOPA. This was shown for monoenzymatic AADC‐neurons of the SCN (D13) (Ishida et al., 2002), premamillary nucleus (D18), pretectal nucleus (D15), and the nucleus of the solitary tract (D2) (Karasawa et al., 1994). Furthermore, the monoenzymatic AADC‐neurons, e.g., the neurons of the dorsomedial nucleus and some hypothalamic neurons in culture become serotonin‐ immunoreactive after the systemic administration of 5‐hydroxytryptophan, but not L‐tryprophan. The preliminary pharmacological inhibition of AADC has abolished this effect thereby confirming the AADC activity in the monoenzymatic neurons (De Vitry et al., 1986). Interestingly, at least a part of monoenzymatic AADC‐neurons lacks the MA transporters. This has been specifically confirmed by the inability of monoenzymatic AADC‐neurons of the SCN for uptake of the radiolabeled serotonin and DA in young and adult rats (Beaudet and Descarries, 1979; Bosler and Calas, 1982; Ugrumov et al., 1986). Although DA synthesis in monoenzymatic AADC‐neurons has been proven, the mechanism of DA storage and release remains uncertain, particularly because of the absence of VMAT2 (Weihe et al., 2006). Nevertheless, DA is probably synthesized in cytosol of monoenzymatic AADC‐neurons as in catecholaminergic neurons. Regarding the DA release, it has been demonstrated ex vivo and in the primary cell culture that under membrane depolarization a large amount of DA is discharged from the AN of rat fetuses (> Figure 2-22) (Melnikova et al., 1998, 1999) containing a large number of monoenzymatic AADC‐ neurons but almost lacking bienzymatic (TH and AADC) neurons (> Figure 2-12) (Ershov et al., 2002a). On one hand these data strongly suggest that the monoenzymatic AADC‐neurons synthesize and store DA, and on the other hand they are capable of releasing DA in response to the adequate physiological stimuli. Still, the source of L‐DOPA for DA synthesis in monoenzymatic AADC‐neurons, mechanism of the DA store and release should be clarified. Although a wide distribution of monoenzymatic AADC‐neurons all over the brain has been repeatedly reported (Jaeger et al., 1984), little is known about their functional significance. In this context, an

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. Figure 2‐22 Dopamine (DA) concentration and Kþ‐stimulated release in: (a) dissected mediobasal hypothalamus (arcuate nucleus & median eminence) of rat fetuses on the 21st day of intrauterine development; (b) primary cell culture of mediobasal hypothalamus of rat fetuses taken on the 17th day of intrauterine development and maintained for 7 days culture (Melnikova et al., 1999). Mean SEM. *P < 0.05

evaluation of intraneuronal colocalization of AADC with neurotransmitters or neuromodulators is of particular importance. Still, only the AADC colocalization with vasopressin in the SCN neurons has been definitely proven by using double‐immunolabeling technique (Jaeger et al., 1983). Nevertheless, a number of other monoenzymatic AADC‐neurons appear to coexpress neuropeptides also. This suggestion is supported by the overlapping in the distribution of monoenzymatic AADC‐neurons and the enkephalin‐producing neurons in the lateral parabrachial nucleus (D3), as well as the substance P‐ and neurotensin‐producing neurons of the nucleus of the solitary tract (D2) (Jaeger et al., 1984). Taking into account that the monoenzymatic AADC‐neurons were often seen in close topographic relations with cerebral blood vessels (Ugrumov et al., 1989a; Karasawa et al., 1994), one may suggest that these neurons synthesize DA and serotonin from the nearest precursors circulating in blood (Melnikova et al., 2006).

4.3 Ensembles of Monoenzymatic Neurons All the previous attempts to evaluate a functional significance of monoenzymatic neurons were restricted to TH‐neurons and AADC‐neurons, separately. Our idea to consider the neurons expressing individual complementary enzymes of DA synthesis as a functional unit occurred to be more productive for understanding of their physiological role. According to our hypothesis, L‐DOPA synthesized in monoenzymatic TH‐neurons is discharged to the intercellular space and thereafter captured by monoenzymatic AADC‐neurons for further conversion to DA (> Figure 2-23) (Ugrumov et al., 2002). Bearing in mind that neurotransmitters and apparently L‐DOPA diffuse for a long distance along the intercellular clefts (Schneider et al., 1994), the cooperative synthesis of DA may be realized even if monoenzymatic TH‐neurons and AADC‐neurons are located far from each other. The AN of fetal rats containing more than 99% monoenzymatic TH‐neurons and AADC‐neurons, in proportion 1:1 and less than 1% bienzymatic neurons has been used as a model to test the hypothesis about the cooperative synthesis of DA by non‐dopaminergic (non‐DA‐ergic) neurons. It has been demonstrated in the ex vivo and in vitro (primary tissue culture) study that despite a minor number of bienzymatic neurons in the AN of fetal rats, the DA concentration in this local region is higher (> Figure 2-22) (Melnikova et al.,

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. Figure 2‐23 Schematic representation of the hypothesis on: (a) cooperative synthesis of dopamine (DA) by monoenzymatic neurons expressing complementary enzymes of DA synthetic pathway (left side); (b) strengthening of DA synthesis in DA‐ergic neurons due to L‐DOPA produced by monoenzymatic tyrosine hydroxylase‐expressing neurons (centre); (c) turning on DA synthesis in serotoninergic neurons due to L‐DOPA produced by monoenzymatic tyrosine hydroxylase‐expressing neurons (right side). AADC, aromatic L‐amino acid decarboxylase; TH, tyrosine hydroxylase; TryH, tryptophan hydroxylase

1999) than in the whole diencephalon or in the whole brain which contains a lot of bienzymatic DA‐ergic neurons (Coyle and Henry, 1973). Furthermore, the L‐DOPA concentration in the AN of fetal rats exceeds three times that of DA (> Figures 2‐20 and > 2‐22) (Melnikova et al., 1999), whereas in the brain regions containing the accumulations of true DA‐ergic neurons, only trace amounts of L‐DOPA are detectable as the enzymatic activity of AADC greatly exceeds that of TH (Moore et al., 1985). Taken together these data have been considered as the indirect indication of DA cooperative synthesis by monoenzymatic neurons. The hypothesis about cooperative synthesis of DA was additionally supported by the observation of close topographic relations between monoenzymatic TH‐neurons and AADC‐neurons in the rat AN that was shown with double‐immunofluorecent labeling of the enzymes in confocal microscopy (Ershov et al., 2002b). Besides simple appositions, sort of specialized‐like junctions were observed. These axo‐somatic contacts were formed by ramified monoenzymatic TH‐axons spreading along a cell body of monoenzymatic AADC‐neurons (> Figure 2-14) (Ershov et al., 2002b). Close topographic relations between monoenzymatic neurons are supposed to serve for increasing an efficacy of the L‐DOPA transfer from the monoenzymatic TH‐neurons to the monoenzymatic AADC‐neurons (> Figure 2-13). The contacts between monoenzymatic TH‐neurons and AADC‐neurons have been observed not only at the level of cell bodies in the AN but also at the level of distal axons including axonal terminals in the external zone of the median eminence (> Figure 2-13). Apart from axo‐axonal contacts, numerous monoenzymatic TH‐axons and AADC‐axons abut on the primary capillary plexus of the hypophysial portal circulation, giving rise to axo‐vascular contacts. The particularly high density of the axo‐vascular contacts of this kind was observed in the lateral region of the median eminence. Despite the great value of double‐immunoflurescent observations in confocal microscopy, the resolution of this technique is not

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sufficient for recognition of true appositions between monoenzymatic neurons. Nevertheless, the very recent electron microscopic study with the double‐immunolabeling of the enzymes of DA synthesis confirmed definitely the existence of direct contacts between monoenzymatic TH‐neurons and AADC‐ neurons (Sorokin et al., unpublished). Despite convincingness of the earlier ex vivo and in vitro data, they may be considered only as indirect evidence of the DA cooperative synthesis in the AN since the input of the minor population of bienzymatic DA‐ergic neurons to DA synthesis cannot be ignored. All the attempts to eliminate completely bienzymatic neurons by using 6‐OHDA were unsuccessful because of the low affinity uptake of DA and hence neurotoxin in this particular brain region (Ershov et al., 2005). The only way to definitely prove the cooperative synthesis of DA by monoenzymatic neurons was to inhibit the L‐DOPA transfer from the monoenzymatic TH‐neurons to monoenzymatic AADC‐neurons. If the hypothesis is valid, this action should provoke a decrease of DA synthesis by monoenzymatic neurons but not by DA‐ergic neurons. In a whole, this should result in a drop of the content and hence synthesis of DA in the AN. Three types of the experimental model could theoretically be used to achieve this result: (1) an inhibition of L‐DOPA release from monoenzymatic TH‐neurons; (2) an anchoring of L‐DOPA in the extracellular space; and (3) an inhibition of the L‐DOPA uptake by the monoenzymatic AADC‐neurons. Finally, the third model has been chosen (> Figure 2-24) (Ugrumov et al., 2004). The presumptive L‐DOPA . Figure 2‐24 Schematic representation (a) and results (b, c) of the experiment showing that the competitive inhibition of neutral amino acid and L‐DOPA transporter with L‐tyrosine (a) resulted in the decrease of dopamine (DA) synthesis in cell suspension of the arcuate nucleus of rat fetuses on the 21st day of prenatal life containing mostly monoenzymatic TH‐neurons and AADC‐neurons (b) and in the increase of DA synthesis in cell suspension of the substantia nigra of the same fetuses containing mostly DA‐ergic neurons (c) (Ugrumov et al., 2004) Tyr (), incubation of cell suspension in the absence of L‐tyrosine; Tyr (þ), incubation of cell suspension in the presence of L‐tyrosine. AADC, aromatic L‐amino acid decarboxylase; TH, tyrosine hydroxylase. Filled oval, neutral amino acid and L‐DOPA transporter

uptake by monoenzymatic AADC‐neurons was inhibited under static or perifusion incubation of the cell suspension of the AN of fetal rats (21st embryonic day) in the presence of a relatively high amount of L‐tyrosine which competes with L‐DOPA for the membrane transporter (> Figure 2-24). The cell suspension of the ventral mesencephalon (substantia nigra) of the same fetuses containing a lot of DA‐ergic

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neurons has been used as a control. The total amount of L‐DOPA or DA in the incubation medium and cell extracts after the incubation has been considered as an index of their synthesis rate. The L‐tyrosine administration resulted in almost 50% increase of the total amount of L‐DOPA in the cell suspensions of the AN and the substantia nigra (Ugrumov et al., 2004). This result was quite predictable as L‐tyrosine stimulates L‐DOPA synthesis in any cell with enzymatically active TH because of the semi‐ saturation of TH with L‐tyrosine under physiological concentration (Dairman, 1972). The influence of L‐tyrosine on DA synthesis in the substantia nigra was opposite in sign to that in the AN. It stimulated significantly DA synthesis in DA‐ergic neurons of the substantia nigra as a substrate of synthesis, and inhibited DA synthesis in the AN as a competitive inhibitor of the L‐DOPA transporter (> Figure 2-24). This result was considered as convincing evidence of cooperative synthesis of DA by monoenzymatic TH‐neurons and AADC‐neurons (Ugrumov et al., 2004). Despite detection of a relatively high level of DA synthesis in the AN of rat fetuses ex vivo and in the cell culture (Melnikova et al., 1999), it remained uncertain whether the DA amount is sufficient to provide the inhibitory control of the adenohypophysial prolactin secretion in fetuses as in adulthood (McCann et al., 1984). The pharmacological model of the inhibition of D2 receptors on the adenohypophysial lactotropes by the systemic administration of haloperidol was used to solve this issue (Melnikova et al., 1998). It has been shown that the inhibitory control of the adenohypophysial prolactin secretion is established in rats during last 2 days of the intrauterine development, almost in the absence of bienzymatic DA‐ergic neurons in the AN. This result is considered as an additional argument in favor of cooperative synthesis of DA by monoenzymatic neurons in the AN (Melnikova et al., 1998). L‐DOPA synthesized in monoenzymatic TH neurons may be involved in cooperative synthesis of DA when captured not only to monoenzymatic AADC‐neurons but also to DA‐ergic and serotoninergic neurons (> Figure 2-23) (Arai et al., 1995; Karasawa et al., 1995; Ugrumov et al., 2002; Kannari et al., 2006). Bearing in mind that the enzymatic activity of AADC greatly exceeds that of TH and tryptophan hydroxylase, the admission of extracellular L‐DOPA to catecholaminergic neurons and serotoninergic neurons promotes catecholamine synthesis or triggers DA synthesis, respectively (> Figure 2-23). Cooperative synthesis of DA by monoenzymatic TH‐neurons and AADC‐neurons as well as by monoenzymatic TH‐neurons and either DA‐ergic neurons or serotoninergic neurons is considered as a compensatory reaction under the failure of DA‐ergic neurons (Ugrumov et al., 2002, 2004). The former takes place in the brain regions containing monoenzymatic TH‐neurons and AADC‐neurons, (e.g., in the AN), whereas the latter might be a characteristic of the brain regions possessing monoenzymatic TH‐neurons and either DA‐ergic neurons or serotoninergic neurons (e.g., the AN and the striatum). In addition to monoenzymatic AADC‐neurons, the glial cells and endothelial cells of blood vessels possess enzymatically active AADC (Hardebo et al., 1980; Juorio et al., 1993). These cells are probably capable of synthesizing DA from L‐DOPA derived either from the monoenzymatic TH‐neurons or from general circulation (Melnikova et al., 2006). It should be emphasized that the penetration of DA synthesized in endothelial cells to the brain is prohibited by the blood–brain barrier.

4.4 Non‐MA‐ergic Neurons Expressing the MA Transporters The neurons partly expressing the serotoninergic phenotype have been first recognized in the eighties in rodents, both in adulthood (Frankfurt and Azmitia, 1983; De Vitry et al., 1986) and in ontogenesis (Ugrumov et al., 1986, 1989a), by using autoradiography and a combination of pharmacological approach and immunocytochemistry for serotonin. Later, this discovery has been confirmed by using the direct technical approaches, double‐labeling of serotonin and the serotonin transporter, and sophisticated animal genetic models (Gaspar et al., 2003). It has been definitely shown that these neurons possess the serotonin transporter but lack tryptophan hydroxylase. However, the expression of AADC remains to be under question (De Vitry et al., 1986). It should be stressed that the neurons partly expressing the serotoninergic phenotype is an attribute of a wide range species, from lobsters to humans (Verney et al., 2002; Richards et al., 2003). The non‐MA‐ergic neurons possessing the mechanism of the serotonin uptake most probably serve to capture and store serotonin released from the next serotoninergic axons arising from the raphe nucleus.

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This conclusion is derived from the observation of the accumulation of serotonin‐immunoreactive material in these neurons following the pharmacological stimulation of serotonin synthesis (L‐tryptophan, 5‐hydroxytryptophan, and pargyline) that may be abolished by the preliminary pharmacological inhibition of the serotonin uptake (Ugrumov et al., 1989a). The earlier suggestion is strongly supported by the fact that the non‐MA‐ergic neurons expressing the serotonin transporter are often located around cerebral ventricles (the dorsomedial nucleus, SCN), close to the supraependymal and subependymal plexes of serotoninergic fibers (Ugrumov et al., 1985b), or in the vicinity to the medial forebrain bundle (lateral preoptic area) composed of serotoninergic and other fibers (> Figure 2-18) (Ugrumov et al., 1989a). There is a certain similarity in functioning of the neurons and platelets which do not synthesize serotonin but capture it from plasma with the serotonin transporter (Maurer‐Spurej, 2005). Non‐MA‐ergic neurons serving to store serotonin most probably compensate the local serotonin deficiency that might be of particular importance in ontogenesis during so‐called critical period of serotonin action on the target‐neurons as a morphogenetic factor (Lauder, 1993; Ugrumov, 1997). If the earlier neurons possess AADC in addition to the serotonin transporter, they may synthesize serotonin from extracellular 5‐hydroxytryptophan including that circulating in blood. Indeed, the neurons abutting on the blood vessels become serotonin‐immunoreactive only after systemic pretreatment of the animals with either pargiline, the MA oxidase inhibitor, or 5‐hydroxytryptophan (Ugrumov et al., 1989a). In this case, serotonin may contribute to the regulation of the vascular tone as a vasoconstrictor (Ugrumov et al., 1989a). Although a number of data in adult mammals may be interpreted in favor of the existence of the non‐ MA‐ergic neurons expressing the DA transporter but lacking either TH or AADC, no direct evidence has been yet obtained (see > Section 3.2). Moreover, no data are available about these neurons in the brain in ontogenesis. Nevertheless, by analogy with the non‐MA‐ergic neurons expressing the serotonin transporter, one can suggest that the non‐MA‐ergic neurons expressing the DA transporter serve for capturing and storage of extracellular DA, as well as for DA synthesis from the extracellular L‐DOPA if expressing AADC additionally. It is important to ascertain in the future study whether the DA accumulated in the non‐DA‐ergic neurons can be further discharged. Indeed, Hoffman et al. (1998), who failed to detect VMAT2 mRNA in neurons expressing DA transporter but lacking TH, have stated that these neurons are capable of removing extracellular DA but may not release it.

5

Tuberoinfundibular Neurons Partly Expressing DA‐ergic Phenotype in Hyperprolactinemia

Tuberoinfundibular system consists of DA‐ergic neurons located in the AN and projecting their axons to the median eminence toward the primary vascular plexus of the hypophysial portal circulation (> Figure 2-25). DA delivered from DA‐ergic axons to the hypophysial portal circulation arrives with the bloodstream to the anterior lobe providing an inhibitory control of prolactin secretion via D2 receptors on lactotropes (McCann et al., 1984). Degeneration of DA‐ergic neurons of the tuberoinfundibular nucleus (¼AN in rodents) in humans leads to the development of the syndrome of hyperprolactinemia and finally to the disturbance of reproduction (Serri et al., 2003). Hyperprolactinemia is reproduced in rats by the injection of 6‐OHDA (neurotoxin) (> Figure 2-26) to the cerebral ventricles (Ershov et al., 2005). One to two weeks after the 6‐OHDA administration, 50–60% DA‐ergic neurons of the AN are degenerated (Ershov et al., 2005) that results in 50% decrease of DA synthesis and in doubling of prolactin concentration in plasma (> Figure 2-26) (Ziyazetdinova et al., unpublished). A relatively low level of the neurotoxin‐induced degeneration of DA‐ergic neurons in the AN compared to DA‐ergic neurons of other location in the brain, e.g., in the substantia nigra (Smith and Helme, 1974; Jonsson, 1983) is explained by the low‐affinity uptake of DA in the AN (Demarest and Moore, 1980; Annunziato et al., 1980;Moore et al., 1985) and by the relatively small number of the neurons expressing the DA transporter (Bosler and Calas, 1982; Hoffman et al., 1998). The prolactin concentration in plasma returns to the normal level in rats 1–1.5 months after the treatment with neurotoxin that is a manifestation of the compensation of the functional insufficiency of

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. Figure 2‐25 Schematic representation of dopaminergic tuberoinfundibular and nigrostriatal systems and their characteristics in norm and pathology

the tuberoinfundibular DA‐ergic system (> Figure 2-26) (Ziyazetdinova et al., unpublished). In theory, the compensatory mechanisms may be represented by: (1) intensified DA synthesis in survived DA‐ergic neurons due to the increased synthesis and/or activity of TH; (2) stimulated cooperative DA synthesis by monoenzymatic neurons due to the increased number of monoenzymatic neurons or the increased activity of the enzymes in preexisting monoenzymatic neurons; (3) enhanced sensibility of lactotropes to DA as a result of the elevated expression of D2 receptors; and (4) attenuated sensibility of lactotropes to stimulating effects of prolactin‐releasing neurohormones like thyrotropin‐releasing hormone and vasoactive intestinal polypeptide. In reality, normalization of the prolactin secretion in the neurotoxin‐treated rats is accompanied with the increased number of the monoenzymatic TH‐neurons and AADC‐neurons (Ershov et al., 2005) and the augmentation of DA synthesis to normal level (Ziyazetdinova et al., unpublished) that was shown both in vivo and in vitro (perifusion of slices of the AN). Taken together these data strongly suggest that the functional insufficiency of the tuberoinfundibular DA‐ergic system is compensated at least in part by the intensified cooperative synthesis of DA by monoenzymatic neurons. Thus, the functional insufficiency of the tuberoinfundibular DA‐ergic system, e.g., in hyperprolactinemia, is compensated due to the stimulation of DA synthesis most probably by the monoenzymatic neurons.

6

Striatal Neurons Partly Expressing DA‐ergic Phenotype in Parkinson’s Disease

Nigrostriatal DA‐ergic system consists of DA‐ergic neurons located in the compact zone of the substantia nigra which project their axons to the striatum (> Figure 2-25). DA released from the distal axons in the striatum plays a key role in the regulation of a motor behavior. That is why the degeneration of nigrostriatal DA‐ergic neurons in humans leads to the disturbance of the motility which is the crucial mechanism of the Parkinson’s disease pathogenesis (Agid, 1991). Noteworthy, clinical symptoms first appear 25–30 years after the onset of the Parkinson’s disease under degeneration of about 70–80% DA‐ergic neurons in the

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. Figure 2‐26 Schematic representation of the experiment (a) and results (b–d) showing the 6‐hydroxydopamine‐induced degeneration of 50% bienzymatic (TH and AADC) neurons in the arcuate nucleus of adult rats is followed first by decreased DA synthesis and increased prolactin secretion (14th day) and then by the normalization of dopamine synthesis and prolactin secretion. Moreover, the number of monoenzymatic neurons increased significantly (Ershov et al., 2005; unpublished data). AN, arcuate nucleus; AADC, aromatic L‐amino acid decarboxylase; TH, tyrosine hydroxylase, 6‐OHDA, 6‐hydroxydopamine

substantia nigra (> Figure 2-27) and the loss of about 70% DA in the striatum (Agid, 1991). This raised a question what mechanisms of the brain plasticity serve to compensate the failure of the nigral DA‐ergic neurons for a long time and what may be the role of the striatal neurons partly expressing the DA‐ergic phenotype in this phenomenon.

6.1 Monoenzymatic TH‐Expressing Neurons The degeneration of the DA‐ergic neurons in the substantia nigra is accompanied by the increased number of the striatal TH‐immunoreactive neurons in all mammals, rodents (Lopez‐Real et al., 2003), monkeys, and humans studied so far (Betarbet et al., 1997; Porritt et al., 2000; Palfi et al., 2002). As observed from the double‐labeling studies, most of them are monoenzymatic (> Figure 2-28) (Lopez‐Real et al., 2003; Sorokin et al., unpublished). In animals, the maximal – 3.5–7‐fold – increase of TH‐immunoreactive neurons after the degeneration of nigral DA‐ergic neurons has been observed in monkeys (> Figure 2-29) (Betarbet et al., 1997; Palfi et al., 2002). In fact, the total number of the TH‐immunoreactive neurons in the denervated

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. Figure 2‐27 Schematic representation of timing characteristics of degeneration of nigral dopaminergic neurons with age in norm and in Parkinson’s disease and its relevance to presymptomatic and symptomatic phases of the disease (adapted from Agid, 1991). Initial number of nigral dopaminergic neurons is considered as 100%

striatum in monkeys greatly exceeded that in rodents (140 neurons per 50 mm‐thick section in monkeys vs. occasional neurons per 40 mm‐thick section in rats). Apparently, this cannot be explained by the species‐ specific difference in the total number of all the striatal neurons only (Betarbet et al., 1997). The decreased number (by sixfold) of the striatal TH‐immunoreactive neurons after degeneration of the nigral DA‐ergic neuron has been described so far in only one paper devoted to the striatum of patients with Parkinson’s disease (Huot et al., 2007). According to the authors’ opinion, the long‐term treatment of these patients with L‐DOPA resulted in a partial compensation of the striatal DA deficiency thereby exerting a hypothetical negative feedback on the TH expression in the striatal neurons. In animals with degenerated nigral DA‐ergic neurons as in intact or control animals, most TH‐immunoreactive neurons were located in the dorsal striatum in the subcallosal area and in the ventral striatum around the anterior commissure (> Figure 2-30) (Meredith et al., 1999; Palfi et al., 2002; Lopez‐ Real et al., 2003). Less numerous TH‐immunoreactive neurons were found scattered in other striatal regions (> Figure 2-30) (Lopez‐Real et al., 2003). Two morphological types of TH‐immunoreactive neurons have been distinguished (Lopez‐Real et al., 2003). Most neurons (99%) of the first type were unipolar, oval in shape and small in size (6–12 mm) having one or two aspiny processes. These were most probably interneurons (Betarbet et al., 1997; Lopez‐Real et al., 2003). Rare neurons ( Figure 2-31) (Betarbet et al., 1997; Lopez‐Real et al., 2003). These neurons were similar in appearance to projection neurons (Lopez‐Real et al., 2003). Some authors emphasized that the TH‐immunoreactive neurons in the denervated striatum were more heavily stained than those in the intact striatum (Betarbet et al., 1997). The 6‐OHDA injection to the substantia nigra in rats resulted in the appearance of new striatal TH‐immunoreactive neurons on the next day and in the progressive increase of their number for two to three subsequent weeks. Then, the TH‐immunoreactive neurons gradually decreased in number for 2–3 months (> Figure 2-32) (Meredith et al., 1999; Lopez‐Real et al., 2003). Similar observations were made in mice after their combined systemic treatment with 1‐methyl‐4‐phenyl‐1,2,3,6‐tetrahydropyridine (MPTP) and 3‐nitropropionic acid. The latter is a semi‐specific neurotoxin providing a general damaging action on the striatal tissue. After the treatment with 6‐OHDA, first TH‐immunoreactive neurons appear in the striatum on the next day after administration of toxins (5.3 neurons per striatum). However, later the number of these neurons first increased rapidly for 3 days reaching maximum on the fourth day (135 neurons per striatum) and then decreased gradually and disappeared by the 80th day (Nakahara et al., 2001).

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. Figure 2‐28 Mono‐ (A, B, E, F) and double‐immunolabeled (C, D) neurons for either tyrosine hydroxylase (A, B, C, E, F) or aromatic L‐amino acid decarboxylase (D) in the rat striatum after 6‐hydroxydopamine‐induced degeneration of nigral dopaminergic neurons (Sorokin et al., unpublished). A‐D, double‐immunofluorescent for tyrosine hydroxylase and aromatic L‐amino acid decarboxylase in confocal microscopy; E, F, mono‐immunolabeling with peroxidase for tyrosine hydroxylase in conventional microscopy. Arrow, immunoreactive neurons. Bar scale: A, B – 10 mm; C, D – 20 mm; E, D – 40 mm

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. Figure 2‐29 Tyrosine hydroxylase‐immunoreactive cell counts in the striatum of control and 1‐methyl‐4‐phenyl‐1,2,3,6‐ tetrahydropyridine(MPTP)‐treated monkeys (Betarbet et al., 1997). *P < 0.05

. Figure 2‐30 Schematic representation of the distribution of tyrosine hydroxylase‐immunoreactive neurons (a) and of aromatic L‐amino acid decarboxylase‐immunoreactive neurons (b) in the rat striatum after 6‐hydroxydopamine‐induced degeneration of nigral dopaminergic neurons (Lopez‐Real et al., 2003). ac, anterior commissure; cc, corpus callosum; v, lateral ventricle

6.2 Monoenzymatic AADC‐Expressing Neurons According to Tashiro et al. (1989b), the number of the striatal AADC‐immunoreactive neurons in rats increases four times under degeneration of the nigral DA‐ergic neurons (12–60 neurons per striatum vs. 3–15 neurons per striatum in the control). Noteworthy, the frequency of the striatal AADC‐immunoreactive neurons appears to be proportional to the damaged area of the compact zone of the substantia nigra. As in intact and control animals, in the neurotoxin‐treated animals most AADC‐immunoreactive neurons are concentrated in the striatum in the subcallosal region, along the lateral ventricle and scattered in other striatal regions (> Figure 2-30) showing a partial overlapping with the location of less numerous

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. Figure 2‐31 Aspiny (a) and spiny (b) tyrosine‐hydroxylase‐immunoreactive neurons in the striatum of 1‐methyl‐4‐phenyl‐ 1,2,3,6‐tetrahydropyridine(MPTP)‐treated monkeys (Betarbet et al., 1997). The insets in a and b show a magnified image of a portion of the dendrite denoted by the arrows

. Figure 2‐32 The number of TH‐immunoreactive neurons at six rostrocaudal levels of the rat striatum (S1–S6) 1, 2, 3, and more than 4 weeks after 6‐hydroxydopamine injection to the raphe nucleus (Meredith et al., 1999)

TH‐immunoreactive neurons (Lopez‐Real et al., 2003). The frequency of the AADC neurons decreased in the rostrocaudal extension of the striatum (Mura et al., 1995, 2000). Striatal AADC‐immunoreactive neurons are small in size (6–10 mm) and oval or spindle‐like in form possessing aspiny processes. They look like interneurons (Mura et al., 1995; Meredith et al., 1999; Lopez‐ Real et al., 2003). According to Meredith et al. (1999), the AADC‐immunoreactive neurons almost disappear with time after the neurotoxin injection, in rats after three weeks.

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6.3 Bienzymatic TH‐ and AADC‐Expressing Neurons It has been confirmed by using the double‐immunolabeling of the enzymes of DA synthesis that the striatum of 6‐OHDA‐treated animals contains bienzymatic neurons (> Figure 2-28) as that in intact (control) animals (Lopez‐Real et al., 2003; Sorokin et al., unpublished). However, the proportion of the striatal monoenzymatic and bienzymatic neurons varied when injecting the neurotoxin to different compartments of the nigrostriatal system. Among the neurons expressing enzymes of DA synthesis, most neurons were bienzymatic after the neurotoxin injection to the striatum whereas their portion did not exceed 30% when the neurotoxin was injected to the medial forebrain bundle (Lopez‐Real et al., 2003). It should be emphasized that in bienzymatic neurons the concentration of the TH‐immunoreactive material was always higher than that of AADC‐immunoreactive material. Therefore, the authors supposed that even a part of the TH‐immunoreactive but AADC‐immunonegative neurons may in fact coexpress both enzymes but AADC is not visible because of its low concentration (Lopez‐Real et al., 2003). The observation of the striatal bienzymatic neurons raised a question whether these neurons are DA‐ergic in nature coexpressing the DA transporter. Although nobody has yet attempted to use triple‐ labeling for solving this issue, it was shown with the double‐immunolabeling technique that in the normal and denervated striatum of the monkeys and humans almost all TH‐immunoreactive neurons coexpress the DA transporter and VMAT2. The number of a such type of the neurons multiplies under degeneration of nigral DA‐ergic neurons (Betarbet et al., 1997; Porritt et al., 2000; Cossette et al., 2005; Tande´ et al., 2006). According to Porritt et al. (2000), the TH‐immunoreactive neurons coexpressing the DA transporter are distributed in the striatum of patients with Parkinson’s disease as follows: 39.9% neurons are located in the putamen, 11.6% in the caudate nucleus, 16.3% in the globus pallidus externa, 6.3% in the globus pallidus interna, and 25.9% in the internal capsule and ansa lenticularis. In addition to the DA transporter, the TH‐immunoreactive neurons coexpress the nuclear orphan receptor Nurr1 (Cossette et al., 2004, Hout and Parent, 2007), a transcription factor essential for the expression of the DA‐ergic phenotype by midbrain neurons (Saucedo‐Cardenas et al., 1998). In addition to bienzymatic neurons expressing the DA transporter and VMAT2, the bienzymatic neurons lacking VMAT2 have been recently found thereby raising the question about DA storage and release in these neurons (Weihe et al., 2006). The same discussion may be addressed to this issue as in the case of monoenzymatic AADC‐neurons (see, > Section 5.2).

6.4 Origin, Functional Properties, and Functional Significance of Striatal Neurons Partly or Completely Expressing the DA‐ergic Phenotype If such a large number of striatal neurons expressing TH and AADC somehow contribute to DA synthesis, this compensatory mechanism may be of a substantial functional importance. Therefore, a number of studies have addressed this issue by reproducing parkinsonism in animals, mostly in rodents and monkeys. The authors attempted to determine the origin, functional significance, and regulation of the neurons expressing enzymes of DA synthesis. The degeneration of DA‐ergic neurons in the substantia nigra in animals was provoked by electrochemical lesion or more often by the administration of 6‐OHDA and MPTP. The latter is administered systemically and transformed in the brain to 1‐methyl‐4‐pyridinium, a specific neurotoxin of DA‐ergic neurons, by MA oxidase B (Nakahara et al., 2001). All the earlier manipulations resulted in the appearance of the striatal neurons possessing enzymes of DA synthesis (Betarbet et al., 1997; Meredith et al., 1999; Palfi et al., 2002; Lopez‐Real et al., 2003). This was shown by using immunocytochemistry, in situ hybridization, the reverse transcriptase reaction followed by polymerase chain reaction (Nakahara et al., 2001). Although the striatal neurons expressing enzymes of DA synthesis were detected in parkinsonian animals most often with mono‐immunolabeling, the major TH‐immunoreactive neurons and AADC‐immunoreactive neurons were supposed to be monoenzymatic as they differ to a certain extent in location, size, and morphology (Meredith et al., 1999). Origin of the striatal neurons partly or completely expressing the DA‐ergic phenotype. Two sources of the striatal neurons partly or completely expressing the DA‐ergic phenotype are considered in literature: the

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stem cells of the lateral ventricles and the preexisting neurons, ‘‘silent’’ in norm and expressing enzymes of DA synthesis and the DA transporter under the local DA depletion. The first hypothesis appears to be nonvaluable as the neurons expressing enzymes of DA synthesis and the DA transporter first appear in the striatum on the next day after the neurotoxin administration, i.e., during the too short period for the origin of neuron from the stem cells, its migration to the striatum, and its differentiation. Furthermore, the cells derived from stem cells and migrated to the striatum do not express specific neuronal markers, NeuN, TH, AADC, or GABA‐decarboxylase even three weeks after their appearance (Nakahara et al., 2001). Therefore, the second hypothesis appears to be more convincing. In fact, the enzymes of DA synthesis are most probably coexpressed in preexisting GABA‐ergic neurons under the local DA deficiency that was shown in the in vivo and in vitro studies by using the double‐immuno‐labeling technique (Max et al., 1996; Betarbet et al., 1997; Mura et al., 2000). Functional properties and functional significance of the striatal neurons partly or completely expressing the DA‐ergic phenotype. Despite the absence of direct evidence of the enzymatic activity of TH in the striatal neurons, most authors consider that TH is capable to convert L‐tyrosine to L‐DOPA. The identification of the principal neurotransmitter or neuromodulator in the striatal neurons coexpressing TH may be useful also for understanding their functional significance. It has been demonstrated both in vivo (Betarbet et al., 1997) and in cell culture (Max et al., 1996) that TH is coexpressed in the GABA‐ergic neurons. In the denervated striatum of monkeys, 99% TH‐immunoreactive neurons are GABA‐ergic and only 1% synthesize calbindin, a marker of striatal projection neurons, or parvalbumin, a marker of a distinct set of striatal interneurons (Betarbet et al., 1997). A substantial number of the TH‐immunoreactive neurons expresses the subunit NR1 of the N‐methyl‐D‐aspartate glutamate receptors (26% TH‐neurons) or the subunit GluR1 of the a‐amino‐3‐hydroxy‐5‐methyl‐isoxazole‐4‐propionic acid glutamate receptors (75% TH‐neurons) though no TH‐immunoreactive neurons express subunits GluR2/3 of the a‐amino‐3‐hydroxy‐5‐methyl‐ isoxazole‐4‐propionic acid type of glutamate receptors or subunits mGluR1/5 of the metabotropic glutamate receptors (Betarbet and Greenamyre, 1999). AADC in monoenzymatic striatal neurons is capable of converting L‐DOPA to DA that was proven by the concomitant appearance of the AADC‐immunoreactive neurons and the DA‐immunoreactive neurons identical by morphology and location in the striatum of the rats following the 6‐OHDA‐induced degeneration of the nigral DA‐ergic neurons (Meredith et al., 1999). Similar DA‐immunoreactive neurons were observed in the striatum following the systemic administration of exogenous L‐DOPA (> Figure 2-33)

. Figure 2‐33 Schematic representation of the distribution of aromatic L‐amino acid decarboxylase‐immunoreactive neurons (a) and dopamine‐immunoreactive neurons (b) in the rat striatum after L‐DOPA administration (Mura et al., 2000). AC, anterior commissure; CC, corpus callosum; LV, lateral ventricle

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(Mura et al., 1995, 2000). Apparently, DA may be synthesized in monoenzymatic AADC‐neurons not only from exogenous L‐DOPA but also from endogenous L‐DOPA circulating with blood (Melnikova et al., 2006) or synthesizing in the striatal monoenzymatic TH‐neurons. As monoenzymatic TH‐neurons, most monoenzymatic AADC‐neurons were GABA‐ergic, and only rare neurons (4.6%) were peptidergic synthesizing calretenin. All the attempts to detect AADC in peptidergic neurons producing somatostatin and parvalbumin were unsuccessful (Cossette et al., 2005b). One cannot exclude that DA produced in monoenzymatic AADC‐neurons interacts functionally with colocalized GABA or calretinin. Apart from monoenzymatic AADC‐neurons and DA‐ergic fibers survived after the neurotoxin administration, endogenous L‐DOPA can be converted to DA in the striatal serotoninergic fibers (Arai et al., 1996; Lopez et al., 2001; Lopez‐Real et al., 2003; Kannari et al., 2006) that is considered as a compensatory mechanism under the local DA deficiency. This suggestion is supported by a number of observations in the denervated striatum: (1) the colocalization of DA in the serotoninergic fibers after systemic administration of L‐DOPA (Maeda et al., 2005); (2) hyperinnervation of the striatum with serotoninergic fibers; and (3) fourfold decrease of the DA extracellular concentration after the additional degeneration of serotoninergic fibers (Tanaka et al., 1999). In addition to serotoninergic fibers, surviving DA‐ergic fibers and monoenzymatic AADC‐neurons, DA is believed to be synthesized in the striatum in the glial cells and endothelial cells containing AADC (Hardebo et al., 1980). However, this suggestion should be taken with caution as: (1) Nakamura et al. (2000) failed to observe the DA‐immunoreactive cells in the primary striatal culture following the L‐DOPA administration; (2) DA synthesized in the endothelial cells probably cannot penetrate to the brain tissue because of the blood–brain barrier. Degeneration of the nigral DA‐ergic neurons is accompanied by the appearance of the neurons expressing enzymes of DA synthesis not only in the striatum but also in adjacent brain regions. Monoenzymatic TH‐immunoreactive neurons appear in the nucleus accumbens and the olfactory bulbs whereas AADC‐neurons become visible in the cortex and in the bed nucleus of the stria terminalis. If these neurons contribute to DA synthesis, it may further diffuse to the striatum providing a volume transmission effect on the target neurons. This is in agreement with the Schneider et al. (1994) data showing that extracellular DA diffuses in the denervated striatum for a particularly long distance (5–7 mm) that is possible due to the loss of DA‐ergic fibers and hence the decrease of the DA uptake (Bergstrom and Garris, 2003; Bezard et al., 2003). The expression of the enzymes of DA synthesis in non‐DA‐ergic neurons outside the striatum probably represents one of the mechanisms of the so‐called ‘‘passive stabilization’’ serving to maintain a normal level of extracellular DA in the denervated striatum without compensatory change of the DA uptake and release (Bergstrom and Garris, 2003). By analogy with hyperprolactinemia following degeneration of the tuberoinfundibular DA‐ergic neurons, one may expect that the DA cooperative synthesis by striatal monoenzymatic neurons is turned on as a compensatory reaction under degeneration of the nigral DA‐ergic neurons in Parkinson’s disease. In fact, the number of the striatal neurons expressing enzymes of DA synthesis in patients with Parkinson’s disease exceeded that of normal humans (Porritt et al., 2000). About 66, 000 neurons expressing either TH or DA transporter have been detected postmortem in the striatum and in the close basal ganglia like globus pallidus and internal capsule in humans with Parkinson’s disease (Porritt et al., 2000). It should be emphasized that this population is only twice as little compared to the whole population of DA‐ergic neurons in the substantia nigra of the same patients and is as large as the population of DA‐ergic neurons of the substantia nigra innervating the putamen in normal humans. Noteworthy, the implantation of only a slightly higher number of embryonic DA‐ergic neurons (80, 000) to patients with Parkinson’s disease is sufficient to induce the substantial but temporal improvement of their status (Ugrumov, 2001). Compensatory mechanisms in Parkinson’s disease. A delay for 25–30 years in the appearance of the initial symptoms of Parkinson’s disease after the onset of the nigral DA‐ergic neuron degeneration is apparently explained by turning on the compensatory processes which are mostly realized in the striatum (Bezard et al., 2003). In addition to the expression of the enzymes of DA synthesis in non‐DA‐ergic neurons, they include: (1) an increase of TH activity and DA synthesis in the surviving nigral DA‐ergic neurons (Agid et al., 1973; Mogi et al., 1988) despite of downregulation of the TH synthesis (Sherman and Moody, 1995); (2) the increased release of DA from the rest of axon terminals (Bernheimer et al., 1973; Zhang et al., 1988); (3) a decrease of the expression of the DA transporter and the DA uptake (Uhl et al.,

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1994; Laihinen et al., 1995; Garris et al., 1997); (4) the enhanced sensibility of the target‐neurons according to upregulation of DA receptors (Bezard et al., 2003); (5) the DA diffusion from intact to denervated regions along the intercellular clefts that is a milestone of the hypothesis on ‘‘the passive stabilization’’ (Bergstrom and Garris, 2003), on one hand, and fits well in the hypothesis on ‘‘the volume or extrasynaptic transmission’’ (Zoli et al., 1998; Vizi, 2000), on the other. The earlier compensatory mechanisms result in maintaining of the normal concentration of DA in denervated striatum under the continuous degeneration of the nigral DA‐ergic neurons (Robinson and Whishaw, 1988; Garris et al., 1997; Bezard et al., 2003). Obviously the list of the compensatory mechanisms is not limited by those described earlier that makes it necessary to extend these studies. After the initial appearance of symptoms, the Parkinson’s disease is developed rapidly as a result of the failure of DA‐ergic system on one hand, and on the other hand by the exhaustion of compensatory resources. Thus, the functional failure of the nigrostriatal DA‐ergic system is temporarily compensated by turning on the mechanisms of the brain plasticity, including the expression of enzymes of DA synthesis in the striatal non‐DA‐ergic neurons which probably contribute to DA synthesis.

7

Regulation of the Partial Expression of MA‐ergic Phenotype by the Brain Neurons in Norm and Pathology

From the earlier data it follows that the frequency of monoenzymatic neurons varies in certain brain regions in ontogenesis and in adulthood under different functional conditions or in pathology. For instance, the number of monoenzymatic neurons decreases in the AN in ontogenesis and increases in adulthood under degeneration of DA‐ergic neurons (Ershov et al., 2002a, 2005). The latter phenomenon is also true for striatal monoenzymatic neurons in adulthood after degeneration of the nigral DA‐ergic neurons (Lopez‐ Real et al., 2003). Another example is represented by magnocellular vasopressinergic neurons of the supraoptic, paraventricular, and accessory nuclei which coexpress TH in norm under chronic functional stimulation (> Figure 2-6) (Abramova et al., 2002), chronic deficiency of vasopressin (Kiss and Mezey, 1986), as well as in pathology under certain metabolic disorders (Fetisov et al., 1997), and perinatal hypoxia (Panayotacopoulou et al., 1994).

7.1 Regulation of the Partial Expression of MA‐ergic Phenotype by Neural Afferents Although the regulation of the expression of the enzymes of DA synthesis in non‐MA‐ergic neurons remains uncertain, some data suggest that neural afferents or more precisely their neurotransmitters contribute to this control. In fact, the number of monoenzymatic neurons increases in certain brain regions, the supraoptic nucleus, the AN and the striatum, after their surgical deafferentation (Kiss and Mezey, 1986; Daikoku et al., 1986; Betarbet et al., 1997; Lopez‐Real et al., 2003) or, on the contrary, decreased in number along with their innervation in ontogenesis (Sorokin et al., unpublished). The earlier hypothesis has been first tested by using vasopressinergic neurons, one of the most promising cell models to solve this problem. Indeed, it has been shown till now that for these cells they may transiently coexpress TH during the osmotic stimulation. This suggests an existence of extracellular signals which can turn on the TH expression after the onset of osmotic stimulation and turn it off after normalization of the water‐mineral metabolism (Yagita et al., 1994). Indeed, it has been recently demonstrated that the TH expression in vasopressinergic neurons is inhibited by noradrenergic afferents and hence noradrenaline via adrenoreceptors (> Figure 2-34). This conclusion derived from the fact that the administration in vivo of the a1‐adrenoreceptor antagonist increased whereas the administration of the a1‐adrenoreceptor agonist decreased the concentrations of TH mRNA and protein in osmotically stimulated vasopressinergic neurons in young rats (Ugrumov, 2002; Abramova et al., unpublished). The catecholaminergic inhibitory control of the expression of the enzymes of MA synthesis in non‐MA‐ ergic neurons may be an attribute not only of vasopressinergic neurons but also of the neurons located

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. Figure 2‐34 Schematic drawing of the inhibitory influence of noradrenergic afferents and noradrenaline on tyrosine hydroxylase expression in vasopressinergic neurons of supraoptic nucleus. dots, noradrenaline; asterisk, tyrosine hydroxylase; filled circles, vasopressin secretory granules; filled triangle, adrenoreceptor

outside the magnocellular nuclei. In this context, the non‐MA‐ergic neurons of the AN, striatum, and the olfactory bulbs appear to be under the DA‐ergic afferent inhibitory control. This suggestion is supported by the observations in rats of: (1) the increased number of monoenzymatic TH‐neurons in the AN (Ershov et al., 2005), striatum (Tashiro et al., 1989a, b), and the olfactory bulbs (Tashiro et al., 1990) under degeneration of their afferent DA‐ergic neurons located in the AN, substantia nigra, and ventral tegmental area, respectively; (2) the coexpression of TH in GABA‐ergic neurons after degeneration of their DA‐ergic afferents (Meredith et al., 1999); (3) the existence of numerous TH‐immunoreactive neurons in the striatum and the limbic system during prenatal and early postnatal periods in rats, before the establishment of their DA‐ergic synaptic innervation, and (4) the disappearance of TH‐immunoreactive neurons by puberty (Sorokin et al., unpublished), after completion of synaptogenesis. The inhibitory action on the TH expression if exists, is provided by DA rather than by L‐DOPA as L‐DOPA does not influence the expression of at least AADC in the striatal neurons (Lopez‐Real et al., 2003). The hypothesis about the catecholaminergic inhibitory control of the enzymes of MA synthesis in non‐ MA‐ergic neurons should be carefully tested in the future because of ambiguity of the interpretations of

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some supporting arguments. In fact, the synchronization of the disappearance of the neurons expressing enzymes of DA synthesis and the establishment of the afferent catecholaminergic synaptic innervation in ontogenesis may be a result not only of the catecholaminergic inhibitory control of the enzymes expression in this local brain area but also of the neuron migration to other brain regions or their apoptosis. Furthermore, the following data also appear to contradict this idea: (1) the monoenzymatic neurons expressing TH are usually located in the denervated striatum in close vicinity of surviving DA‐ergic fibers (Porritt et al., 2000; Lopez‐Real et al., 2003); (2) DA promotes the stimulating influence of some neurotropic factors, e.g., ‘‘brain‐derived neurotrophic factor’’, on TH expression in the primary cell culture of the striatum (Du and Iacovitti, 1995).

7.2 Paracrine Regulation of the Partial Expression of MA‐ergic Phenotype by Diffusive Factors Neurotrophic or growth factors which are produced to a great extent by glial cells appear to be among the most efficient chemical signals stimulating the expression of the enzymes of DA synthesis in non‐MA‐ergic neurons. This suggestion has derived from the observation of the synchronization of the increased secretion of neurotrophic factors (Nakajima et al., 2001) with the expression of the enzymes of DA synthesis in the striatal non‐DA‐ergic neurons under degeneration of nigral DA‐ergic neurons. The intensity of both processes increased continuously until reaching maximum 2–3 weeks following the neurotoxin injection (Meredith et al., 1999; Nakajima et al., 2001). The neurotrophic factors were shown to stimulate the surviving of DA‐ergic neurons and the local reparative processes in the denervated striatum of adult animals, as well as in the striatum of ageing animals (Du and Iacovitti, 1995; Tomac et al., 1995). They occur to be of particular efficiency in the brain of primates. This is manifested by the multiple increase of the number of the neurons expressing TH and the DA transporter in the intact striatum of old monkeys or in the DA‐ergic denervated striatum in young monkeys after the intracerebral injections of the vector of the glia‐derived neurotrophic factor to both groups of the animals (Palfi et al., 2002). A number of data obtained in the in vivo and in vitro studies have definitely proven that the neurotrophic factors promoted the expression of the enzymes of DA synthesis in the striatal neurons. The fibroblast growth factor and the brain‐derived neurotrophic factor occurred to be particularly efficient in stimulation of the TH expression in non‐DA‐ergic neurons in the denervated striatum, and their action is mediated by DA. Although the glial‐derived neurotrophic factor and the ciliary neurotrophic factor provide similar action on the striatal neurons, they are less efficient (Du and Iacovitti, 1995). The administration of neurotrophic factors to the striatal tissue culture induced the appearance of the initial TH‐immunoreactive cells in 12 hours. Then, TH‐immunoreactive neurons increased rapidly in number reaching maximum 18 hours after the treatment that was followed by the gradual decrease in the neuron number for subsequent 4 days (Du and Iacovitti, 1995). The action of neurotrophic factors has been shown to be mediated via specific receptors. The glia‐derived neurotrophic factor provides its action through a multireceptor complex composed of a novel glycosylphosphatidylinositol‐anchored glia‐derived neurotrophic factor receptor‐a and the receptor tyrosine kinase product of the c‐ret proto‐oncogene (Durbec et al., 1996; Trupp et al., 1997). The expression of this receptor in the developing brain in rats is maximal in the early postnatal period that coincides with a high level of the TH expression (Sorokin et al., unpublished) and minimal in adulthood (Trupp et al., 1997). In addition to catecholamines (see earlier), vasopressin is also considered as an extracellular signal providing an inhibitory control of the TH expression in vasopressinergic neurons. This was indirectly confirmed by the decreased level of TH in the neurons of the magnocellular nuclei in the vasopressin deficient Brattleboro rats treated with exogenous vasopressin (Kiss and Mezey, 1986). A list of chemical signals controlling the expression of the enzymes of DA synthesis in non‐DA‐ergic neurons apparently is not limited to catecholamines, DA, vasopressin, and neurotrophic factors. By contrast with these extracellular signals, some signals may provide the opposite effects when acting on the neurons in different brain regions or in the same region but over different periods of ontogenesis. For instance, the serotonin deficiency provoked by p‐chlorophenylalanine, an inhibitor of serotonin synthesis, resulted in

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the increased number of TH‐immunoreactive neurons in the dorsal motor vagal nucleus (Kitahama et al., 1987) and in the decrease of the TH content in the differentiating neurons of the AN that was shown both in vivo and in the primary cell culture (Melnikova et al., 2001). Noteworthy, serotonin as a morphogenetic diffusive factor provides a long‐lasting effect on differentiating TH‐neurons in the AN that is manifested by the maintaining of a relatively low level of TH in adult offspring treated with p‐chlorophenylalanine during a certain period of the intrauterine development (Melnikova et al., 2001). The only weak point of the earlier studies was the use of the TH mono‐immunolabeling as it is impossible to precise whether the monoenzymatic TH‐neurons or bienzymatic DA‐ergic neurons are the targets for serotonin. Only occasional studies have been somehow related to the regulation of the expression of AADC in monoenzymatic neurons. It has been demonstrated in rats that the AADC gene expression and AADC activity in monoenzymatic neurons of the SCN strongly depend on the circadian rhythms being maximal during daylight hours and minimal over nocturnal period. Although the regulation of AADC expression in these neurons by extracellular signals is quite probable, their nature remains uncertain (Ishida et al., 2002).

7.3 Hormonal Regulation of the Partial Expression of MA‐ergic Phenotype The regulation of the expression of the enzymes of DA synthesis in non‐MA‐ergic neurons is provided probably not only by the brain‐derived factors but also by hormones. However, no direct evidence of this suggestion is so far available. The most promising model to solve this issue in the future appears to be the AN containing a great portion of monoenzymatic TH‐neurons and AADC‐neurons (Ershov et al., 2002a, b), which synthesize DA in cooperation (Ugrumov et al., 2004), in addition to DA‐ergic bienzymatic neurons . It has been repeatedly demonstrated that the DA‐producing neurons of the AN contribute to the regulation of reproduction providing the inhibitory control of the pituitary prolactin secretion (McCann et al., 1984; Moore et al., 1985). Moreover, DA produced in the AN contributes to the regulation of the gonadotropin‐releasing hormone secretion by the neurons of the anterior forebrain. In turn, DA synthesis in the AN is under the feedback control of the pituitary (prolactin) and gonadal (estrogens, progesterone) reproductive hormones (Arbogast and Voogt, 1991). The hormonal regulation of the DA‐producing neurons in the AN is supported by the observation of receptors for prolactin (Arbogast and Voogt, 1997; Lerant and Freeman, 1998), estrogens (Jones and Naftolin, 1990; Hou et al., 2003;Mitchell et al., 2003), and progesterone (Warembourg et al., 1993; Dufourny et al., 2005) on TH‐expressing neurons, as well as by detection of the progesterone receptors on AADC‐containing neurons (Warembourg et al., 1993). The earlier data were considered as the proof of the receptor expressions in DA‐ergic neurons (Jones and Naftolin, 1990; Arbogast and Voogt, 1991, 1997; Lerant and Freeman, 1998; Hou et al., 2003) though the authors did not attempt to detect both enzymes of DA synthesis in these neurons by using double‐labeling technique. Therefore, it remains uncertain whether the hormone receptors are expressed in DA‐ergic and/or in monoenzymatic neurons. Up to the present, only occasional studies have provided indirect evidence of the expression of receptors for hormones of reproduction both in the DA‐ergic and monoenzymatic neurons. In fact, the number of monoenzymatic TH‐neurons and AADC‐neurons increased significantly under hyperprolactinemia provoked by 6‐OHDA‐induced degeneration of DA‐ergic neurons (Ershov et al., 2005). Apparently, this compensatory mechanism is responsible for normalization with time of the prolactin secretion (Fenske and Wuttke, 1976). The decrease of the density of monoenzymatic TH‐fibers in the median eminence synchronous to a change of the concentrations of prolactin and progesterone in plasma in the rats during postnatal period (Arbogast and Voogt, 1991; Ershov et al., 2002b) may be considered as another argument in favor of the expression of the receptors for reproduction hormones in monoenzymatic TH‐neurons. Furthermore, the data showing that the TH‐expressing neurons located in the dorsomedial area of the AN, mostly bienzymatic, and those located in the ventrolateral area of the AN, mostly monoenzymatic, are differently regulated by reproduction hormones (Lerant and Freeman, 1998) appear to be particularly promising for further strengthening of our knowledge about the hormonal regulation of the

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partial expression of the MA‐ergic phenotype. Noteworthy, the sensitivity to reproduction hormones, e.g., prolactin is an attribute not only of DA‐producing neurons of the tuberoinfundibular system but also of those in other brain regions including the nigrostriatal system (Chen and Ramirez, 1989). Thus, the expression of the enzymes of DA synthesis in monoenzymatic non‐DA‐ergic neurons appears to be under the control of classical neurotransmitters, paracrine diffusive factors, and hormones.

Acknowledgments This study was supported by the grants of: the program of the Presidium of the Russian Academy of Sciences ‘‘Basic sciences for medicine’’, the program of the Department of Biological Sciences ‘‘Physiological mechanisms of the regulation of homeostasis in the systemic control of the animal behavior’’, RGNF 06‐06‐ 000‐10A, Scientific Schools‐6352.2006.4., RFBR 05-04-48829, RFBR-OFI 07-04-12211, PICS 07-04-92173.

References Abramova M, Calas A, Thibault J, Ugrumov M. 2000. Tyrosine hydroxylase in vasopressinergic axons of the pituitary posterior lobe of rats under salt‐loading as a manifestation of neurochemical plasticity. Neural Plast 7: 179-191. Abramova M, Marsais F, Calas A, Thibault J, Ugrumov M. 2002. Dynamical study of tyrosine hydroxylase expression and its correlation with vasopressin turnover in the magnocellular neurons of the supraoptico‐posthypophysial system under long‐term salt‐loading of adult rats. Brain Res 925: 67-75. Agid Y. 1991. Parkinson’s disease pathophysiology. Lancet 337: 1321-1323. Agid Y, Javoy F, Glowinski J. 1973. Hyperactivity of remaining dopaminergic neurons after partial destruction of the nigrostriatal dopaminergic system in the rat. Nat New Biol 245: 150-151. Annunziato L, Leblanc P, Kordon C, Weiner RI. 1980. Differences in the kinetics of dopamine uptake in synaptosome preparations of the median eminence relative to other dopaminergically innervated brain regions. Neuroendocrinology 31: 316-320. Arai R, Karasawa N, Geffard M, Nagatsu I. 1995. L‐DOPA is converted to dopamine in serotonergic fibers of the striatum of the rat: A double‐labeling immunofluorescence study. Neurosci Lett 11: 195-198. Arai R, Karasawa M, Nagatsu I. 1996. Aromatic L‐amino acid decarboxylase is present in serotonergic fibers of the striatum of the rat. A double‐labeling immunofluorescence study. Brain Res 706: 177-179. Arbogast LA, Voogt JL. 1991. Hyperprolactinemia increases and hypoprolactinemia decreases tyrosine hydroxylase messenger ribonucleic acid levels in the arcuate nuclei, but not the substantia nigra or zona incerta. Endocrinology 128: 997-1005.

Arbogast LA, Voogt JL. 1997. Prolactin (PRL) receptors are colocalized in dopaminergic neurons in fetal hypothalamic cell cultures: Effect of PRL on tyrosine hydroxylase activity. Endocrinology 138: 3016-3023. Balan IS, Ugrumov MV, Calas A, Mailly P, Krieger M, et al. 2000. Tyrosine hydroxylase‐ and/or aromatic L‐amino acid decarboxylase‐expressing neurons in the mediobasal hypothalamus of perinatal rats: Differentiation and sexual dimorphism. J Comp Neurol 425: 167-176. Battaglia AA, Beltramo M, Thibault J, Krieger M, Calas A. 1995. A confocal approach to the morphofunctional characterization of the transient tyrosine hydroxylase system in the rat suprachiasmatic nucleus. Brain Res 696: 7-14. Beaudet A, Descarries L. 1979. Radioautographic characterization of a serotonin‐accumulating nerve cell group in the adult rat hypothalamus. Brain Res 160: 231-243. Beltramo M, Calas A, Chernigovskaya N, Borisova N, Polenova O, et al. 1994. Postnatal development of the suprachiasmatic nucleus in the rat. Morphofunctional characteristics and time course of tyrosine hydroxylase immunopositive fibers. Neuroscience 63: 603-610. Berger B, Verney C, Gaspar P, Febvret A. 1985. Transient expression of tyrosine hydroxylase immunoreactivity in some neurons of the rat neocortex during postnatal development. Brain Res 355: 141-144. Bergstrom BP, Garris PA. 2003. ‘‘Passive stabilization’’ of striatal extracellular dopamine across the lesion spectrum encompassing the presymptomatic phase of Parkinson’s disease: A voltammetric study in the 6‐OHDA‐lesioned rat. J Neurochem 87: 1224-1236. Bernheimer H, Birkmayer W, Hornykiewcz O, Jellinger K. Seitelberger R. 1973. Brain dopamine and the syndromes of Parkinson and Huntington. J Neurol Sci 20: 415-455.

Brain neurons partly expressing monoaminergic phenotype Betarbet R, Greenamyre JT. 1999. Differential expression of glutamate receptors by the dopaminergic neurons of the primate striatum. Exp Neurol 159: 401-408. Betarbet R, Turner R, Chockkan V, DeLong MR, Allers KA, et al. 1997. Dopaminergic neurons intrinsic to the primate striatum. J Neurosci 17: 6761-6768. Bezard E, Gross ChE, Brotchie JM. 2003. Presymptomatic compensation in Parkinson’s disease is not dopamine‐ mediated. Trends Neurosci 26: 215-221. Bjo¨rklund A, Lindvall O. 1984. Dopamine‐containing system in the CNS. Handbook of Chemical Neuroanatomy, Vol. 2., Bjo¨rklund A, Ho¨kfelt T, editors. Amsterdam: Elsevier; pp. 55-122. Bosler O, Calas A. 1982. Radioautographic investigation of monoaminergic neurons: An evaluation. Brain Res Bull 9: 151-169. Bosse R, Fumagalli F, Jaber M, Giros B, Gainetdinov RR, et al. 1997. Anterior pituitary hypoplasia and dwarfism in mice lacking the dopamine transporter. Neuron 19: 127-138. Chaillou E, Tramu G, Thibault J, Tillet Y. 1998. Presence of galanin in dopaminergic neurons of the sheep infundibular nucleus: A double staining immunohistochemical study. J Chem Neuroanat 15: 251-259. Chen JC, Ramirez VD. 1989. Effects of prolactin on tyrosine hydroxylase activity of central dopaminergic neurons of male rats. Eur J Pharmacol 166: 473-479. Cossette M, Parent A, Levesque D. 2004. Tyrosine hydroxylasepositive neurons intrinsic to the human striatum express the transcription factor Nurr1. Eur J Neurosci 20: 20892095. Cossette M, Lecomte F, Parent A. 2005a. Morphology and distribution of dopaminergic neurons intrinsic to the human striatum. J Chem Neuroanat 29: 1-11. Cossette M, Le´vesque D, Parent A. 2005b. Neurochemical characterization of dopaminergic neurons in human striatum. Parkinsonism Relat Disord 11: 277-286. Coyle JT, Henry D. 1973. Catecholamines in fetal and newborn rat brain. J Neurochemistry 21: 61-67. Dahlstro¨m A, Fuxe K. 1964. Evidence for the existence of monoamine‐containing neurons in the central nervous system. I. Demonstration of monoamines in the cell bodies of brainstem neurons. Acta Physiol Scand Suppl 232: 1-55. Daikoku S, Kawano H, Okamura I, Tokuzen M, Nagatsu I. 1986. Ontogenesis of immunoreactive tyrosine hydroxylase‐ containing neurons in rat hypothalamus. Brain Res 393: 85-98. Dairman W. 1972. Catecholamine concentrations and the activity of tyrosine hydroxylase after an increase in the concentration of tyrosine in rat tissues. Br J Pharmacol 44: 307-310.

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Demarest KT, Moore KE. 1980. Accumulation of L‐DOPA in the median eminence: An index of tuberoinfundibular dopaminergic nerve activity. Endocrinology 106: 463-468. De Vitry F, Hamon M, Catelon J, Dubois M, Thibault J. 1986. Serotonin initiates and autoamplifies its own synthesis during mouse central nervous system development. Proc Natl Acad Sci USA 83: 8629-8633. Du X, Iacovitti L. 1995. Synergy between growth factors and transmitters required for catecholamine differentiation in brain neurons. J Neurosci 15: 5420-5427. Dubach M, Schmidt R, Kunkel D, Bowden DM, Martin R, et al. 1987. Primate neostriatal neurons containing tyrosine hydroxylase: Immunohistochemical evidence. Neurosci Lett 75: 205-210. Dufourny L, Caraty A, Clarke IJ, Robinson JE, Skinner DC. 2005. Progesterone‐receptive dopaminergic and neuropeptide Y neurons project from the arcuate nucleus to gonadotropin‐releasing hormone‐rich regions of the ovine preoptic area. Neuroendocrinology 82: 21-31. Durbec P, Marcos‐Gutierrez CV, Kilkenny C, Grigoriou M, Suvanto P, et al. 1996. GDNF signaling through the Ret receptor tyrosine kinase. Nature 381: 789-793. Ershov PV, Ugrumov MV, Calas A, Krieger M, Thibault J. 2002a. Differentiation of tyrosine hydroxylase‐expressing and/or aromatic L‐amino acid decarboxylase‐expressing neurons in the rat mediobasal hypothalamus: Quantitative double‐immunofluorescence study. J Comp Neurol 446: 114-122. Ershov PV, Ugrumov MV, Calas A, Krieger M, Thibault J. 2005. Degeneration of dopaminergic neurons triggers an expression of individual enzymes of dopamine synthesis in non‐dopaminergic neurons of the arcuate nucleus in adult rats. J Chem Neuroanat 30: 27-33. Ershov PV, Ugrumov MV, Calas A, Makarenko IG, Krieger M, et al. 2002b. Neurons possessing enzymes of dopamine synthesis in the mediobasal hypothalamus of rats. Topographic relations and axonal projections to the median eminence in ontogenesis. J Chem Neuroanat 24: 95-107. Everitt B, Meister B, Ho¨kfelt T, Melander T, Terenins L, et al. 1986. The hypothalamic arcuate nucleus‐median eminence complex: Immunocytochemistry of transmitters, peptides and DARPP‐32 with special reference to coexistence in dopamine neurons. Brain Res Rev 11: 97-155. Fenske M, Wuttke W. 1976. Effects of intraventricular 6‐hydroxydopamine injections on serum prolactin and LH levels: Absence of stress‐induced pituitary prolactin release. Brain Res 104: 63-70. Ferna´ndez E, Torrents D, Zorzano A, Palacı´n M, Chillaro´n J. 2005. Identification and functional characterization of a novel low affinity aromatic‐preferring amino acid transporter (arpAT). One of the few proteins silenced during primate evolution. J Biol Chem 280: 19364-19372.

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2

Brain neurons partly expressing monoaminergic phenotype

Fetisov S, Marsais F, Nicolandis S, Calas A. 1997. Expression of tyrosine hydroxylase in magnocellular hypothalamic neurons of obese (fa/fa) and lean heterozygous (Fa/fa) Zucker rats. Mol Brain Res 50: 314-318. Fisher A, Biggs CS, Eradiri O, Starr MS. 2000. Dual effects of L‐3,4‐Dihydroxyphenylalanine on aromatic L‐amino acid decarboxylase, dopamine release and motor stimulation in the reserpine‐treated rat: Evidence that behaviour is dopamine dependent. Neuroscience 95: 97-111. Frankfurt H, Azmitia E. 1983. The effect of intracerebral injections of 5,7‐dihydroxytryptamine and 6‐hydroxydopamine on the serotonin‐immunoreactive cell bodies and fibers in the adult rat hypothalamus. Brain Res 261: 91-99. Frankfurt H, Lauder J, Azmitia E. 1981. The immunocytochemical localization of serotoninergic neurons in the rat hypothalamus. Neurosci Lett 24: 227-232. Fuxe K, Ungerstedt U. 1968. Histochemical studies on the effect of (positive)‐amphetamine, drugs of the imipramine group and tryptamine on central catecholamine and 5‐hydroxytryptamine neurons after intraventricular injection of catecholamines and 5‐hydroxytryptamine. Eur J Pharmacol 4: 135-144. Garris PA, Walker QD, Wightman RM. 1997. Dopamine release and uptake rates both decrease in the partially denervated striatum in proportion to the loss of dopamine terminals. Brain Res 753: 225-234. Gaspar P, Berger B, Febvret A, Vigny A, Krieger‐Poulet M, et al. 1987. Tyrosine hydroxylase‐immunoreactive neurons in the human cerebral cortex: A novel catecholaminergic group? Neurosci Lett 80: 257-262. Gaspar P, Cases O, Maroteaux L. 2003. The developmental role of serotonin: News from mouse molecular genetics. Nat Rev Neurosci 4: 1002-1012. Goshima Y, Kubo T, Misu Y. 1988. Transmitter‐like release of endogenous 3,4‐dihydroxyphenylalanine from rat striatal slices. J Neurochem 50: 1725-1730. Goshima Y, Nakamura S, Ohno K, Misu Y. 1991. Picomolar concentrations of L‐dopa stereoselectively potentiate activities of presynaptic b‐adrenoreceptors to facilitate the release of endogenous noradrenaline from rat hypothalamic slices. Neurosci Lett 129: 214-216. Hardebo JE, Emson PC, Falck B, Owman Ch, Rosengren E. 1980. Enzymes related to monoamine transmitter metabolism in brain microvessels. J Neurochem 35: 1388-1393. Hoffman BJ, Hansson SR, Mezey E´, Palkovits M. 1998. Localization and dynamic regulation of biogenic amine transporters in the mammalian central nervous system. Front Neuroendocrinol 19: 187-231. Ho¨kfelt T, Johansson O, Goldstein M. 1984. Central catecholamine neurons as revealed by immunocytochemistry with special reference to adrenaline neurons. Handbook of

Chemical Neuroanatomy, Vol. 2., Bjo¨rklund A, Ho¨kfelt T, editors. Amsterdam: Elsevier; pp. 157-276. Hou Y, Yang SP, Voogt JL. 2003. Changes in estrogen receptor‐a expression in hypothalamic dopaminergic neurons during proestrous prolactin surge. Endocrine 20: 131-138. Huot P, Le´vesque M, Parent A. 2007. The fate of striatal dopaminergic neurons in Parkinson’s disease and Huntington’s chorea. Brain 130: 222-232. Huot P, Parent A. 2007. Dopaminergic neurons intrinsic to the striatum. J Neurochem 101:1441-1447. Ibata Y, Fukui K, Obata HL, Tanaka M, Hisa Y, et al. 1982. Postnatal ontogeny of catecholamine and somatostatin neuron systems in the median eminence of the rat as revealed by a colocalization technique. Brain Res Bull 9: 407-415. Ikemoto K, Kitahama K, Jouvet A, Arai R, Nishimura A, et al. 1997. Demonstration of L‐dopa decarboxylating neurons specific to human striatum. Neurosci Lett 232: 111-114. Ikemoto K, Kitahama K, Nishimura A, Jouvet A, Nishi K, et al. 1999. Tyrosine hydroxylase and aromatic L‐amino acid decarboxylase do not coexist in neurons in the human anterior cingulate cortex. Neurosci Lett 269: 37-40. Ikemoto K, Nagatsu I, Kitahama K, Jouvet A, Nishimura A, et al. 1998a. A dopamine‐synthesizing cell group demonstrated in the human basal forebrain by dual labeling immunohistochemical technique of tyrosine hydroxylase and aromatic L‐amino acid decarboxylase. Neurosci Lett 243: 129-132. Ikemoto K, Nagatsu I, Nishimura A, Nishi K, Arai R. 1998b. Do all of human midbrain tyrosine hydroxylase neurons synthesize dopamine? Brain Res 805: 255-258. Ikemoto K, Nishimura A, Oda T, Nagatsu I, Nishi K. 2003. Number of striatal D‐neurons is reduced in autopsy brains of schizophrenics. Legal Medicine 5: 221-224. Ishida Y, Yokoyama C, Inatomi T, Yagita K, Dong X, et al. 2002. Circadian rhythm of aromatic L‐amino acid decarboxylase in the rat suprachiasmatic nucleus: Gene expression and decarboxylating activity in clock oscillating cells. Genes Cells 7: 447-459. Izvolskaia M, Duittoz AH, Ugrumov M, Tillet Y. 2006. Tyrosine hydroxylase expression in the olfactory/respiratory epithelium in early sheep fetuses (Ovis aries). Brain Res 1083: 29-38. Jaeger CB, Albert VR, Joh TH, Reis DJ. 1983. Aromatic L‐amino acid decarboxylase in the rat brain: Coexistence with vasopressin in small neurons of the suprachiasmatic nucleus. Brain Res 276: 362-366. Jaeger CB, Ruggiero DA, Albert VR, Park DH, Joh TH, et al. 1984. Immunocytochemical localization of aromatic‐ L‐amino acid decarboxylase. Handbook of Chemical

Brain neurons partly expressing monoaminergic phenotype Neuroanatomy, Vol. 2., Bjo¨rklund A, Ho¨kfelt T, editors. Amsterdam: Elsevier; pp 387-408. Jaeger CB, Teitelman G. 1992. Immunocytochemical distribution of aromatic L‐amino acid decarboxylase (AADC) in rat embryos. Handbook of Chemical Neuroanatomy, Vol. 10: Ontogeny of Transmitters and Peptides in the CNS. Bjo¨rklund A, Ho¨kfelt T, Tohyama M, editors. Amsterdam: Elsevier. Jones EE, Naftolin F. 1990. Estrogen effects on the tuberoinfundibular dopaminergic system in the female rat brain. Brain Res 510: 84-91. Jonsson G. 1983. Chemical lesions techniques: Monoamine neurotoxins. Handbook of Chemical Neuroanatomy, Vol. 1: Methods in Chemical Neuroanatomy. Bjo¨rklund A, Ho¨kfelt T, editors. Amsterdam: Elsevier; Pages 463-507. Juorio AV, Li X‐M, Walz W, Paterson IA. 1993. Decarboxylation of L‐dopa by cultured mouse astrocytes. Brain Res 626: 306-309. Kannari K, Huo Shen H, Arai A, Tomiyama M, Baba M. 2006. Reuptake of L‐DOPA‐derived extracellular dopamine in the striatum with dopaminergic denervation via serotonin transporters. Neurosci Lett 402: 62-65. Karasawa N, Arai R, Isomura G, Nagatsu T, Nagatsu I. 1995. Chemical features of monoaminergic and non‐ monoaminergic neurons in the brain of laboratory shrew (Suncus murinus) are changed by systemic administration of monoamine precursors. Neurosci Res 24: 67-74. Karasawa N, Arai R, Isomura G, Nagatsu T, Nagatsu I. 1997. Coexistence of tyrosine hydroxylase and serotonin in the raphe nucleus of the laboratory shrew (Suncus murinus) during postnatal life. Brain Res Dev Brain Res 99: 121-125. Karasawa N, Arai R, Isomura G, Yamada K, Sakai K, et al. 1994. Phenotypic changes of AADC‐only immunopositive premamillary neurons in the brain of laboratory shrew Suncus murinus by systemic administration of monoamine precursors. Neurosci Lett 179: 65-70. Kiss JZ, Mezey E. 1986. Tyrosine hydroxylase in magnocellular neurosecretory neurons. Response to physiological manipulations. Neuroendocrinology 43: 519-525. Kitahama K, Be´rod A, Denoyer M, Jouvet M. 1987. Visualization of tyrosine hydroxylase‐immunoreactive neurons in the cat dorsal motor vagal cells after treatment with parachlorophenylalanine. Neurosci Lett 77: 155-160. Kitahama K, Ikemoto K, Jouvet A, Nagatsu I, Sakamoto N, et al. 1998. Aromatic L‐amino acid decarboxylase‐ and tyrosine hydroxylase‐immunohistochemistry in the adult human hypothalamus. J Chem Neuroanat 16: 43-55. Klein DC, Moore RY, Reppert SM. 1991. Suprachiasmatic Nucleus. The Mind’s Clock. New York: Oxford University Press.

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Komori K, Fujii T, Nagatsu I. 1991. Do some tyrosine hydroxylase‐immunoreactive neurons in the human ventrolateral arcuate nucleus and globus pallidus produce only L‐dopa? Neurosci Lett 133: 203-206. Laihinen AO, Rinne JO, Na˚gren KA˚, Lehikoinen PK, Oikonen VJ, et al. 1995. PET studies on brain monoamine transporters with carbon‐11‐b‐CIT in Parkinson’s disease. J Nucl Med 36: 1263-1267. Lauder J M. 1993. Neurotransmitters as growth regulatory signals: Role of receptors and second messengers. Trends Neurosci 16: 233-240. Lebrand C, Cases O, Wehrle’ R, Blakely RD, Edwards RH, et al. 1998. Transient developmental expression of monoamine transporters in the rodent forebrain. J Comp Neurol 401: 506-524. Lerant A, Freeman ME. 1998. Ovarian steroids differentially regulate the expression of PRL‐R in neuroendocrine dopaminergic neuron populations: A double label confocal microscopic study. Brain Res 802: 141-154. Lopez A, Mun˜oz A, Guerra MJ, Labandeira‐Garcia JL. 2001. Mechanisms of the effects of exogenous levodopa on the dopamine‐denervated striatum. Neuroscience 103: 639-651. Lopez‐Real A, Rodriguez‐Pallares J, Guerra M.J, Labandeira‐ Garcia JL. 2003. Localization and functional significance of striatal neurons immunoreactive to aromatic L‐amino acid decarboxylase or tyrosine hydroxylase in rat Parkinsonian models. Brain Res 969: 135-146. Lorang D, Amara SG, Simerly RB. 1994. Cell‐type‐specific expression of catecholamine transporters in the rat brain. J Neurosci 14: 4903-4914. Maeda T, Nagata K, Yoshida Y, Kannan K. 2005. Serotonergic hyperinnervation into the dopaminergic denervated striatum compensates for dopamine conversion from exogenously administered L‐DOPA. Brain Res 1046: 230-233. Marsais F, Parmentier C, Terao E, Taxi J, Calas A. 2002. Expression of tyrosine hydroxylase and vasopressin in magnocellular neurons of salt‐loaded aged rats. Microsc Res Tech 56: 81-91. Maurer‐Spurej E. 2005. Serotonin reuptake inhibitors and cardiovascular diseases: A platelet connection. Cell Mol Life Sci 62: 159-170. Max SR, Bossio A, Iacovitti L. 1996. Co‐expression of tyrosine hydroxylase and glutamic acid decarboxylase in dopamine differentiation factor‐treated striatal neurons in culture. Brain Res Dev Brain Res 91: 140-142. McCann SM, Lumpkin MD, Mizunuma H, Khorram O, Ottlecz J, et al. 1984. Peptidergic and dopaminergic control of prolactin release. Trends Neurosci 7: 127-131. Meister B, Ho¨kfelt T, Steinbusch HWM, Skagerberg G, Lindvall O, et al. 1988. Do tyrosine hydroxylase‐immunoreactive neurons in the ventro‐lateral arcuate nucleus

69

70

2

Brain neurons partly expressing monoaminergic phenotype

produce dopamine or only L‐DOPA? J Chem Neuroanat 1: 59-64. Melnikova V, Orosco M, Calas A, Sapronova A, Gainetdinov R, et al. 1999. Dopamine turnover in the mediobasal hypothalamus in rat fetuses. Neuroscience 89: 235-241. Melnikova V, Orosco M, Rouch C, Calas A, Nicolaidis S, et al. 1998. Prolactin secretion and its dopamine inhibitory control in rat fetuses. Eur J Endocrinol 139: 337-342. Melnikova VI, Sapronova AY, Lavrentieva AV, Proshlyakova EV, Voronova SN, et al. 2006. Brain is on of the sources of L‐3,4‐dihydroxyphenylalanine in general circulation of fetal and neonatal rats (Russia). J Evol Physiol Biochem 42: 19-25. Melnikova VI, Ugrumov MV, Proshlyakova EV, Calas A, Thibault J. 2001. Tyrosine hydroxylase expression in differentiating neurons of the rat arcuate nucleus: Stimulatory influence of serotonin afferents. Neural Plast 8: 271-284. Meredith GE, Farrell T, Kellaghan P, Tan Y, Zahm DS, et al. 1999. Immunocytochemical characterization of catecholaminergic neurons in the rat striatum following dopamine‐ depleting lesions. Eur J Neurosci 11: 3585-3596. Mirochnik VV, Makarenko IG, Ugryumov MV. 2002. The source of transitory innervation of suprachiasmatic nucleus by tyrosine hydroxylase‐immunoreactive fibers during postnatal period in rats (Russia). Ontogenez 33: 182-186. Misu Y, Kitahama K, Goshima Y. 2003. L‐3,4‐Dihydroxyphenylalanine as a neurotransmitter candidate in the central nervous system. Pharmacol Ther 97: 117-137. Mitchell V, Loyens A, Spergel DJ, Flactif M, Poulain PM, et al. 2003. A confocal microscopic study of gonadotropin‐ releasing hormone (GnRH) neuron inputs to dopaminergic neurons containing estrogen receptor alpha in the arcuate nucleus of GnRH‐green fluorescent protein transgenic mice. Neuroendocrinology 77: 198-207. Mogi M, Harada M, Kiuchi K, Kojima K, Kondo T, et al. 1988. Homospecific activity (activity per enzyme protein) of tyrosine hydroxylase increases in Parkinson’s brain. J Neural Transm 72: 77-81. Mons N, Tison F, Geffard M. 1989. Identification of L‐dopa‐ dopamine and L‐dopa cell bodies in the rat mesencephalic dopaminergic cell systems. Synapse 4: 99-105. Montange M, Calas A. 1988. Serotonin and endocrinology – the pituitary. Neural Serotonin. Osborne NN, Hamo M, editors. Chichester: John Wiley & Sons; pp. 271-303. Moore KE, Riegle GD, Demarest KT. 1985. Regulation of tuberoinfundibular dopaminergic neurons: Prolactin and inhibitory neuronal influences. Catecholamines as Hormone Regulators. Raven Press; New York: pp. 31-48. Mura A, Jackson D, Manley MS, Young SJ, Groves PM. 1995. Aromatic L‐amino acid decarboxylase immunoreactive cells

in rat striatum: A possible site for the conversion of exogenous L‐DOPA to dopamine. Brain Res 704: 51-60. Mura A, Linder JC, Young SJ, Groves PM. 2000. Striatal cells containing aromatic L‐amino acid decarboxylase: An immunohistochemical comparison with other classes of striatal neurons. Neuroscience 98: 501-511. Nagatsu T. 1995. Tyrosine hydroxylase: Human isoforms, structure and regulation in physiology and pathology. Essays Biochem 30: 15-35. Nagatsu I, Komori K, Takeuchi T, Sakai M, Yamada K, et al. 1990. Transient tyrosine hydroxylase‐immunoreactive neurons in the region of the anterior olfactory nucleus of pre‐ and postnatal mice do not contain dopamine. Brain Res 511: 55-62. Nagatsu I, Sakai M, Takeuchi T, Arai R, Karasawa N, et al. 1997. Tyrosine hydroxylase (TH)‐only‐immunoreactive non‐catecholaminergic neurons in the brain of wild mice or the human TH transgenic mice do not contain GTP cyclohydrolase I. Neurosci Lett 228: 55-57. Nagatsu I, Takeuchi T, Sakai M, Karasawa N, Yamawaki Y, et al. 1996. Transient appearance of tyrosine hydroxylase‐ immunoreactive non‐catecholaminergic neurons in the medial geniculate nucleus of postnatal mice. Neurosci Lett 211: 183-186. Nakahara T, Yamamoto T, Endo K, Kayama H. 2001. Neuronal ectopic expression of tyrosine hydroxylase in the mouse striatum by combind administration of 1‐methyl‐4‐phenyl‐ 1,2,3,6‐tetrahydropyridine and 3‐nitropropionic acid. Neuroscience 108: 601-610. Nakajima K, Hida H, Shimano Y, Fujimoto I, Hashitani T, et al. 2001. GDNF is a major component of trophic activity in DA‐depleted striatum for survival and neurite extension of DAergic neurons. Brain Res 916: 76-84. Nakamura K, Ahmed M, Barr E, Leiden JM, Kang UJ. 2000. The localization and functional contribution of striatal aromatic L‐amino acid decarboxylase to L‐3,4‐dihydroxyphenylalanine decarboxylation in rodent parkinsonian models. Cell Transplant 9: 567-576. Nakamura S, Goshima Y, Yue JL, Misu Y. 1992. Transmitter‐ like basal and K(þ)‐evoked release of 3,4‐dihydroxyphenylalanine from the striatum in conscious rats studied by microdialysis. J Neurochem 58: 270-275. Novak CM, Nunez AA. 1998. Tyrosine hydroxylase‐ and/or aromatic L‐amino acid decarboxylase‐containing cells in the suprachiasmatic nucleus of the Syeian hamster (Mesocricetus auratus). J Chem Neuroanat 14: 87-94. Okamura H, Kitahama K, Mons N, Ibata Y, Jouver M, et al. 1988a. L‐DOPA‐immunoreactive neurons in the rat hypothalamic tuberal region. Neurosci Lett 95: 42-46. Okamura H, Kitahama K, Nagatsu I, Geffard M. 1988b. Comparative topography of dopamine‐ and tyrosine

Brain neurons partly expressing monoaminergic phenotype hydroxylase‐immunoreactive neurons in the rat arcuate nucleus. Neurosci Lett 95: 347-353. Okamura H, Kitahama K, Raynaud B, Nagatsu I, Borri‐ Volttatorni C, et al. 1988c. Aromatic L‐amino acid decarboxylase (AADC)‐immunoreactive cells in the tuberal region of the rat hypothalamus. Biomed Res 9: 261-267. Okamura H, Murakami S, Chihara K, Nagatsu I, Ibata Y. 1985. Coexistence of growth hormone releasing factor‐like and tyrosine hydroxylase‐like immunoreactivities in neurons of the rat arcuate nucleus. Neuroendocrinology 41: 177-179. Palfi S, Leventhal L, Chu Y, Ma SY, Emborg M, et al. 2002. Lentivirally delivered glial cell line‐derived neurotrophic factor increases the number of striatal dopaminergic neurons in primate models of nigrostriatal degeneration. J Neurosci 22: 4942-4954. Panayotacopoulou MT, Raadsheer FC, Swaab DF. 1994. Colocalization of tyrosine hydroxylase with oxytocin or vasopressin in neurons of the human paraventricular and supraoptic nucleus. Brain Res Dev Brain Res 83: 59-66. Porritt MJ, Batchelor PE, Hughes AJ, Kalnins R, Donnan GA, et al. 2000. New dopaminergic neurons in Parkinson’s disease striatum. Lancet 356: 44-45. Richards KS, Simon DJ, Pulver SR, Beltz BS, Marder E. 2003. Serotonin in the developing stomatogastric system of the lobster, Homarus americanus. J Neurobiol 54: 380-392. Robinson TE, Whishaw IQ. 1988. Normalization of extracellular dopamine in striatum following recovery from a partial unilateral 6‐OHDA lesion of the substantia nigra: A microdialysis study in freely moving rats. Brain Res 450: 209-244. Sakai M, Fujii T, Karasawa N, Arai R, Nagatsu I. 1995. Enhanced expression of tyrosine hydroxylase and aromatic L‐amino acid decarboxylase in cerebellar Purkinje cells of mouse after hyperosmotic stimuli. Neurosci Lett 194: 142-144. Saucedo‐Cardenas O, Quintana‐Hau JD, Le WD, Smidt MP, Cox JJ, et al. 1998. Nurr1 is essential for the induction of the dopaminergic phenotype and the survival of ventral mesencephalic late dopaminergic precursor neurons. Proc Natl Acad Sci USA 95: 4013-4018. Schneider JS, Rothblat DS, DiStefano L. 1994. Volume transmission of dopamine over large distances may contribute to recovery from experimental Parkinsonism. Brain Res 643: 86-91. Serri O, Chik CL, Ur E, Ezzat S. 2003. Diagnosis and management of hyperprolactinemia. CMAJ 169: 575-581. Sherman TG, Moody CA. 1995. Alterations in tyrosine hydroxylase expression following partial lesions of the nigrostriatal bundle. Brain Res Mol Brain Res 29: 285-296.

2

Skagerberg G, Meister B, Ho¨kfelt T, Lindvall O, Goldstein M, et al. 1988. Studies on dopamine‐, tyrosine hydroxylase‐ and aromatic L‐amino acid decarboxylase‐containing cells in the rat diencephalon: Comparison between formaldehyde‐induced histofluorescence and immunofluorescence. Neuroscience 24: 605-620. Smith GC, Helme RD. 1974. Ultrastructural and fluorescence histochemical studies on the effects of 6‐hydroxydopamine on the rat median eminence. Cell Tissue Res 152: 493-512. Squire LR, Bloom FE, McConnell SK, Roberts JL, Spitzer NC, et al. 2003. Fundamental Neuroscience. Amsterdam: Academic Press. Steinbusch HWM. 1984. Serotonin‐immunoreactive neurons and their projections. Handbook of Chemical Neuroanatomy, Vol. 3., Classical CNS Transmitters and Transmitter receptors in the CNS. II. Bjo¨rklund A, Ho¨kfelt T, Kuhar MJ, editors. Amsterdam: Elsevier; pp. 68-125. Sugaya Y, Sasaski Y, Goshima Y, Kitahama K, Kusakabe T, et al. 2001. Autoradiographic studies using L‐(14C)DOPA and 3 þ L‐( H)DOPA reveal regional Na ‐dependent uptake of the neurotransmitter candidate L‐DOPA in the CNS. Neuroscience 104: 1-14. Tanaka H, Kannari K, Maeda T, Tomiyama M, Suda T, et al. 1999. Role of serotonergic neurons in L‐DOPA‐derived extracellular dopamine in the striatum of 6‐OHDA‐ lesioned rats. Neuroreport 1: 631-634. Tande´ D, Ho¨glinger G, Debeir T, Freundlieb N, Hirsch EC, et al. 2006. New striatal dopamine neurons in MPTP‐ treated macaques result from a phenotypic shift and not neurogenesis. Brain 129: 1194-2000. Tashiro Y, Kaneko T, Nagatsu I, Kikuchi H, Mizuno N. 1990. Increase of tyrosine hydroxylase‐like immunoreactive neurons in the nucleus accumbens and the olfactory bulb in the rat with the lesion in the ventral tegmental area of the midbrain. Brain Res 531: 159-166. Tashiro Y, Kaneko T, Sugimoto T, Nagatsu I, Kikuchi H, et al. 1989a. Striatal neurons with aromatic L‐amino acid decarboxylase‐like immunoreactivity in the rat. Neurosci Lett 100: 29-34. Tashiro Y, Sugimoto T, Hattori T, Uemura Y, Nagatsu I, et al. 1989b. Tyrosine hydroxylase‐like immunoreactive neurons in the striatum of the rat. Neurosci Lett 97: 6-10. Tinner B, Fuxe K, Ko¨hler C, Hersh L, Andersson K, et al. 1989. Evidence for the existence of a population of arcuate neurons costoring choline acetyltransferase and tyrosine hydroxylase immunoreactivities in the male rat. Neurosci Lett 99: 44-49. Tomac A, Lindqvist E, Lin LF, Ogren SO, Young D, et al. 1995. Protection and repair of the nigrostriatal dopaminergic system by GDNF in vivo. Nature 373: 335-339.

71

72

2

Brain neurons partly expressing monoaminergic phenotype

Trupp M, Belluardo N, Funaksohi H, Ibanez CF. 1997. Complementary and overlapping expression of glial cell line‐derived neurotrophic factor (GDNF), c‐ret proto‐ oncogene, and GDNF receptor‐a indicates multiple mechanisms of trophic actions in the adult rat CNS. J Neurosci 17: 3554-3567. Ugrumov MV. 1997. Hypothalamic monoaminergic systems in ontogenesis: Development and functional significance. Int J Dev Biol 41: 809-816. Ugrumov MV. 2001. Neurotransplantation in treatment of Parkinson’s disease. Basal Ganglia and Thalamus in Health and Movement Disorders. Kultas‐Ilinsky K, Ilinsky IA, editors. New York: Kluwer Academic Publishers/Plenum; pp. 349-363. Ugrumov MV. 2002. Magnocellular vasopressin system in ontogenesis: Development and regulation. Microsc Res Tech 56: 164-171. Ugrumov MV, Ivanova IP, Mitskevich MS, Liposits Z, Setalo, et al. 1985a. Axovascular relationships in developing median eminence of perinatal rats with special reference to luteinizing hormone‐releasing hormone projections. Neuroscience 16: 897-906. Ugrumov M, Melnikova V, Ershov P, Balan I, Calas A. 2002. Tyrosine hydroxylase‐ and/or aromatic L‐amino acid decarboxylase‐expressing neurons in the rat arcuate nucleus: Ontogenesis and functional significance. Psychoneuroendocrinology 7: 533-548. Ugrumov MV, Melnikova VI, Lavrentyeva AV, Kudrin VS, Rayevsky KS. 2004. Dopamine synthesis by non‐dopaminergic neurons expressing individual complementary enzymes of the dopamine synthetic pathway in the arcuate nucleus of fetal rats. Neuroscience 124: 629-635. Ugrumov MV, Popov AP, Vladimirov SV, Kasmambetova S, Thibault J. 1994. Development of the suprachiasmatic nucleus in rats during ontogenesis: Tyrosine hydroxylase immunopositive cell bodies and fibers. Neuroscience 58: 151-160. Ugrumov MV, Taxi J, Mitskevich MS, Arluison M, Tramu G. 1985b. Immunocytochemical and radioautographic study of serotonin projections to cerebral ventricles of perinatal rats. Brain Res 350: 225-230. Ugrumov MV, Taxi J, Mitskevich MS, Tramu G. 1986. Development of the hypothalamic serotoninergic system during ontogenesis in rats. Immunocytochemical and radioautographic study. Brain Res 395: 75-84. Ugrumov MV, Taxi J, Steinbusch HWM, Tramu G, Mitskevich MS. 1989a. On the distribution and morpho‐functional characteristics of 5‐HT‐immunoreactive cells in the hypothalamus of fetuses and neonatal rats. Brain Res Dev Brain Res 46: 233-241

Ugrumov M, Taxi J, Tixier‐Vidal A, Thibault J, Mitskevich MS. 1989b. Ontogenesis of tyrosine hydroxylase‐immunopositive structures in the rat hypothalamus. An atlas of neuronal cell bodies. Neuroscience 29: 135-156. Ugrumov M, Taxi J, Tixier‐Vidal A, Thibault J, Mitskevich MS. 1989c. Ontogenesis of tyrosine hydroxylase‐immunopositive structures in the rat hypothalamus. Fiber pathways and terminal fields. Neuroscience 29: 157-166. Uhl GR, Walther D, Mash D, Faucheux B, Javoy‐Agid F. 1994. Dopamine transporter messenger RNA in Parkinson’s disease and control substantia nigra neurons. Ann Neurol 35: 494-498. Van den Pol AN, Herbst RS, Powell JF. 1984. Tyrosine hydroxylase‐immunoreactive neurons of the hypothalamus: A light and electron microscopic study. Neuroscience 13: 1117-1156. Verney C, Gaspar P, Febvret A, Berger B. 1988. Transient tyrosine hydroxylase‐like immunoreactive neurons contain somatostatin and substance P in the developing amygdala and bed nucleus of the stria terminalis of the rat. Brain Res 470: 45-58. Verney C, El Amraoui A, Zecevic N. 1996. Comigration of tyrosine hydroxylase and gonadotropin‐releasing hormone‐ immunoreactive neurons in the nasal area of human embryos. Brain Res Dev Brain Res 97: 251-259. Verney C, Lebrand C, Gaspar P. 2002. Changing distribution of monoaminergic markers in the developing human cerebral cortex with special emphasis on the serotonin transporter. Anat Rec 267: 87-93. Vincent SR, Hope BT. 1990. Tyrosine hydroxylase containing neurons lacking aromatic amino acid decarboxylase in the hamster brain. J Comp Neurol 295: 290-298. Vizi ES. 2000. Role of high‐affinity receptors and membrane transporters in nonsynaptic communication and drug action in the central nervous system. Pharmacol Rev 52: 63-89. Warembourg M, Varoqueaux F, Poulain P. 1993. Le re´cepteur de la progeste´rone dans les neurones hypothalamiques du cobayae: Colocalisation avec la TH et projections des neurones vers l’e´minence me´deiane. Ann Endocrinol 54: 34 Weihe E, Depboylu C, Schu¨tz B, Scha¨fer MKH, Eiden LE. 2006. Three types of tyrosine hydroxylase‐positive CNS neurons distinguished by dopa decarboxylase and VMAT2 co‐expression. Cell Mol Neurobiol 26: 659-678. Weihe E, Eiden LE. 2000. Chemical neuroanatomy of the vesicular amine transporters. FASEB J 14: 2435-2449. Weiner RI, Findell PR, Kordon C. 1988. Role of classic and peptide neuromediators in the neuroendocrine regulation of LH and prolactin. The Physiology of Reproduction. Knobil E, Neill J, et al. editors. New York: Raven Press; pp. 1235-1281.

Brain neurons partly expressing monoaminergic phenotype Yagita K, Okamura H, Ibata Y. 1994. Rehydration process from salt‐loading: Recovery of vasopressin and its coexisting galanin, dynorphin and tyroisine hydroxylase immunoreactivities in the supraoptic and paraventricular nuclei. Brain Res 667: 13-23. Zhang WQ, Tilson HA, Nanry KP, Hudson PM, Hong JS, et al. 1988. Increased dopamine release from striata of rats after unilateral nigrostriatal bundle damage. Brain Res 461: 335-342.

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Zoli M, Agnati LF, Tinner B, Steinbusch H, Fuxe K. 1993. Distribution of dopamine‐immunoreactive neurons and their relationship to transmitter and hypothalamic hormone‐immunoreactive neuronal systems in the rat mediobasal hypothalamus. A morphometric and microdensitometric analysis. J Chem Neuroanat 6: 293-310. Zoli M, Torri C, Ferrari R, Jansson A, Zini I, et al. 1998. The emergence of the volume transmission concept. Brain Res Brain Res Rev 26: 136-147.

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In Vivo Imaging of Neurotransmitter Systems with PET

B. Gulya´s . C. Halldin . B. Mazie`re

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Introduction: Neurotransmitter and Neuroreceptor Systems and In Vivo Neuroimaging . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 76

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Positron Emission Tomography . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 77

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Labeling Tracers and Ligands with PET Bioisotopes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 79

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Preliminary Steps in the Development of Radioligands for Human CNS Receptors . . . . . . . . . . . . 82

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Measuring Radioligand Effects in the Brain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 83

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Modeling Ligand Effects in the Brain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 85

7 Two Approaches . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 85 7.1 Direct Approach: Radiolabeling of Ligands . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 86 7.2 Indirect Approach: Using Radiolabeled Ligands and Drug Candidates . . . . . . . . . . . . . . . . . . . . . . . . . . . . 86 8 8.1 8.2 8.3 8.4 8.5 8.6 8.7 8.8 8.9

Radioligands for Mapping Neurotransmitter Systems: Some Examples . . . . . . . . . . . . . . . . . . . . . . . . . 87 Dopamine Receptor and Transporter Ligands . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 89 Serotoninergic Neurotransmission Radioligands . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 91 Norepinephrine Neurotransmission Radioligands . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 93 Acetylcholine Radioligands . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 93 Central Benzodiazepine‐Binding Site Ligands . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 93 Peripheral Benzodiazepine Receptor or Benzodiazepine‐Binding Site Receptor Ligands . . . . . . . . . . 94 Glutamate Neurotransmission Radioligands . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 94 Cannabinoid Neurotransmission Radioligands . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 95 Opioid Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 95

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Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 95

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2008 Springer ScienceþBusiness Media, LLC.

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In vivo imaging of neurotransmitter systems with PET

Abstract: The advent of functional neuroimaging techniques has significantly widened the methodological repertoire of neurochemistry. Using positron emission tomography PET, the human and non-human primate brain’s neurotransmitter and neuroreceptor systems can be studied in vivo. With the help of PET the distribution of the various neurotransmitter and neuroreceptor systems can be localized in precise anatomical context and several parameters of these systems can be measured in a quantitative manner. The basics of the technique, development of radiolabelled ligands, modeling and measuring radioligand effects in the brain are, among others, those key issues that are discussed concisely in the present chapter. List of Abbreviations: CT, Computed tomography; FDG, fluoro-deoxy-glucose; HPLC, high performance liquid chromatography; MRI, magnetic resonance imaging; NMSP, N‐methylspiperone; PBBS, peripheral benzodiazepine‐binding site; PET, positron emission tomography; SA, specific activity; SPECT, single‐ photon‐emission computed tomography; SR, specific radioactivity

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Introduction: Neurotransmitter and Neuroreceptor Systems and In Vivo Neuroimaging

In addition to ex vivo–in vitro biochemical probes and postmortem brain imaging techniques, functional neuroimaging techniques, including single‐photon‐emission computed tomography or SPECT and positron emission tomography or PET, have in recent decades become inevitable parts of the methodological armory for the exploration of the human and nonhuman primate brain’s neurotransmitter and neuroreceptor systems. The present chapter focuses on PET research. And as the recent advances in PET radiochemistry result in the emergence of newer and newer PET radioligands with an increasing pace, in the present chapter the major emphasis is on the basic principles of PET radioligand development, and no emphasis is laid on a comprehensive summary of all available radioligands. For this reason, in the first part of this chapter a somewhat detailed overview on the bases of PET neuroreceptor research is given, whereas in the second part of the chapter we provide the reader with a short survey of those established PET radioligands, which have gained application grounds in a number of PET laboratories. During the past decades, well over a hundred different neurotransmitter subsystems have been identified and described in detail. In parallel with this, radiochemistry research has focused on the development of labeled PET radioligands to cover as wide as possible a range of biological markers usable for in vivo brain mapping with PET. As several neurotransmitter and receptor systems have been demonstrated to be directly involved in neurological or psychiatric diseases, drug addiction, or personality disorders, the importance of PET in receptor‐system‐related drug research has increased tremendously in recent years. Take an example! > Figure 3-1 is a schematic overview of the presynaptic generation and synaptic transmission of one of the key monoamine neurotransmitters, dopamine. Dopamine, similarly to several other central neurotransmitters, in addition to its extrasynaptic effects, predominantly acts as a synaptic transmitter (and this is displayed in the figure). Until now, two major families and five subtypes of the dopamine receptor have been identified (the dopamine D1 and D5 receptors in one family, and the D2, D3, and D4 receptors in the other). Most of these receptor subtypes can be targeted with PET radioligands. These receptors are, at the same time, the major targets of neuropsychiatric drugs used e.g., in schizophrenia or other psychiatric diseases. In addition to the receptor systems, the transporter molecule, responsible for the reuptake of the intrasynaptic dopamine molecules, is also a major target of neuropsychiatric drug research, since this molecule is predominantly responsible for the regulation of the intrasynaptic dopamine concentration. Consequently, the transporter system can also be a prime target for PET radioligand development. As exemplified by the figure, in vivo neuroimaging with PET using labeled ligands or ‘‘radioligands’’ can ‘‘visualize’’ the various receptor and transporter systems and measure in quantitative terms their densities, binding and occupancy status, and other parameters. Naturally, for the visualization of these systems other imaging techniques, for example postmortem autoradiography, can also be used. Consequently, the application of radioligands can also help us understand the role of various neurotransmitter systems and their concerted behavior (‘‘receptor fingerprint’’) in the normal and pathological functioning of the human brain. Furthermore, PET imaging of central neuroreceptor systems can greatly contribute to our efforts in developing novel drugs targeting dedicated receptor sites in the human brain.

In vivo imaging of neurotransmitter systems with PET . Figure 3-1 A schematic overview of synaptic dopamine neurotransmission. A number of relevant 3 H‐ or 125I‐labeled autoradiography radioligands are indicated in the figure

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C‐labeled PET and

Positron Emission Tomography

PET is a noninvasive biological imaging technique that can quantitatively measure the distribution of radiolabeled molecules in the living body. Although the spatial resolution of the technique is lagging behind that of dedicated anatomical imaging techniques, in combination with anatomical/morphological imaging methods (CT or MRI), PET can map ‘‘quantitatively’’ biochemical and physiological parameters in a proper anatomical context. On the other hand, the temporal resolution of the method allows us to follow the biodynamics of the labeled molecules in the brain and body. A PET laboratory consists of expensive and sophisticated instrumentation and its activities require concerted actions from a number of experts of various disciplines (> Figure 3-2). The basis of the technique is the detectability of g photons, generated during the annihilation of positrons and electrons (> Figure 3-3). Positron‐emitting radionuclides, including those which are most commonly used in PET (> Table 3-1), decay by emitting a positively charged electron, a positron. The positron is the counterpart of the electron: its mass is identical with that of the electron, but it has a positive charge. Within a short distance (1–2 mm) from the place of the decay, the emitted positron encounters an electron. During the encounter, the two annihilate each other by releasing two 511 keV g photons in opposite directions along an axis. These two g photons can be detected by a pair of scintillation crystals (e.g., NaI or BiGe crystals), which transform the g photons into photons in the range of the visible light spectrum. These scintillation photons are amplified in photomultipliers, and the resulting signals can be detected by a coincidence circuit (> Figure 3-3). In the PET scanner, the detectors are built into detector rings (> Figure 3-4a). Inside a detector ring, with the help of the functional logic of coincidence circuits, one detector can be ‘‘functionally coupled’’ to several other detectors, i.e., a large number of detector channels can be formed (> Figure 3-4b). Several

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. Figure 3-2 The components and main functions of a complex PET center

. Figure 3-3 The physical basis of positron emission and positron–electron annihilation, and coincidence detection with the help of a functionally coupled detector system

. Table 3-1 Positron‐emitting radionuclides commonly used in PET Nuclide T½ (min) Target Nuclear reaction Radioactive decay Maximal energy (MeV) Specific radioactivity (Ci/mmol) Effective dose equivalent per 100 MBq radiotracer (mSv) Common forms Critical organ

11

C 20.3 NþO 14 N(p,a)!11C 11 11 0 6C ! 5B þ þ1e 0.97 9
106 0.4

13

N 9.98 H2O 12 C(d,n)!13N

15

O 2.05 NþO 14 N(d,n)!15O

18

1.20 19
106 0.25

1.74 90
106 0.1

0.64 1.7
106 2.5

11

13

N, 13NH3 Liver

15

O2, C15O Lung

H18F, 18F2 Bladder

CO, 11CO2 Liver

F 110 H218O 18 O(p,n)!18F

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. Figure 3-4 (a) Detector rings inside the scanner. (b) The possible arrangements of detector channels, related to one given detector, in a detector ring. (c) A PET scanner

rings can form the gantry of the scanner so that a larger section of the body can fully be covered by the field of view of the system (> Figure 3-4c). The detected radioactivity distributions inside the body can be reconstructed with the help of filtered backprojection algorithms (> Figure 3-5). Using additional physiological information and mathematical models, from the radioactivity distribution maps, obtained by the PETscanner, a large number of biochemical parameters can be estimated (> Table 3-2).

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Labeling Tracers and Ligands with PET Bioisotopes

Among the PET radionuclides, the frequently utilized 18F has earlier been used in one single form: FDG. More recently, a burgeoning variety of radioligands is available with this radionuclide. The use of 13N and 15 O is more limited, mainly owing to the shorter half‐times of these radionuclides. 15O‐Labeled water or butanol is used in blood flow studies, whereas 13N‐labeled ammonia can be used in cardiac PET studies. Definitely, the most widely used PET radionuclide is 11C. For its ‘‘comfortable’’ half‐time (20 min) and carbon’s central role in organic chemistry, 11C has for long been used in the widest variety of PET radiochemistry applications. An important prerequisite for radioligand development is that the molecule maintains its properties after labeling. This is one further reason for the common use of the short‐lived

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. Figure 3-5 Schematic diagram showing the procedure of filtered backprojection. (a) The original biological target has a radioactivity distribution pattern inside the scanner’s gantry. (b) The main question of data sampling and reconstruction centers around the reliability of the generated image. (c) In the special case of a point source centered in the gantry, all projections, regardless of angle, will be the same. (d) If these projections were to be backprojected, the reconstructed image would be smeared. (e) Instead, a convolution between the profile and the reconstruction filter is applied, creating a filtered profile. (f) The filtered profile can then be used to successfully reconstruct the original image

. Table 3-2 Physiological–biochemical parameters measurable with PET Blood flow Blood volume Protein synthesis Molecular diffusion Tissue pH Metabolism of oxygen, glucose, amino acids, fatty acids, fluor, etc. Receptor and transporter systems: uptake, distribution, binding, occupancy Pharmacodynamics and pharmacokinetics of labeled drugs

positron‐emitting radionuclide 11C, for the substitution of naturally occurring 12C with 11C does not change the biochemistry or the pharmacology of the ligand molecule. The range and number of the applications of 11C‐labeled receptor ligands and drugs have steadily been increasing. The most widely used approach for labeling ligands using 11C is [11C]methylation. A typical total synthesis time for a 11C‐labeled radioligand including HPLC purification is 30 min. The reaction may require the presence of base for the generation of the nucleophile. Moreover, [11C]acylations with [11C]acyl

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chlorides and [11C]cyanation with [11C]cyanide are two common types of reactions, but, for instance, [11C] ethylation and other operations can also be used in [11C] radiochemistry. Nowadays, 18F is used more and more often for labeling central nervous system (CNS) ligands. The unique advantage of 18F lies in its relatively longer half‐time: 110 min, which provides us with the opportunity of transporting the labeled ligands to larger distances, i.e., distributing [18F]‐labeled radioligands to PET scanners not adjacent to a cyclotron facility. The production and application of PET radioligands, used for human studies, requires several consecutive steps, the timing of which is of vital importance due to the decay of the radionuclide and the loss of specific activity (SA) of the labeled radioligand (> Figure 3-6). A general rule of thumb is that the radiochemical synthesis time should start without a delay after the production of the radionuclide and it should be completed within 4–5 half‐life times of the radionuclide, whereas the end of batch efficacy

. Figure 3-6 The flowchart of radioligand production and its use in PET measurements

(radioactivity in the end‐product versus initial radioactivity upon the start of the radiochemistry process) should be over 20%. As in most cases, the radioligand’s SA (the proportion of labeled versus unlabeled molecules in the batch) is an essential parameter and the highest possible SA values are favored; the radioligand should be administered without any significant delay. These rules are less strict in the case of 18 F ligands, as compared with 11C ligands. (Fowler et al. (1999); Guly´as et al. (1998); Haldin et al. (1991, 2001, 2004); Heiss et al. (2001)).

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Preliminary Steps in the Development of Radioligands for Human CNS Receptors

When a potential candidate ligand has been identified and a radiochemical labeling technique developed, some preclinical evaluation, before PET in humans, needs to be performed. In the first instance, useful information may be obtained by studies in rodents. Typically, the radioligand is injected intravenously into a series of rodents. These are then sacrificed at known times after injection and either the animal’s different organs are removed and radioactivity is counted, thus providing the distribution of the radiotracer in different organs over time (tissue distribution study), or the body is sliced, the slices are mounted on glass, and exposed to radiosensitive film (ex vivo autoradiography). In addition to this, clearance of radioactivity from plasma and information on the appearance of labeled metabolites in plasma can be obtained. Specificity of binding may be demonstrated by using selective and potent unlabeled compounds that act in a competing way at the site of interest. A complementary tool in early receptor radioligand development may be autoradiography experiments, wherein frozen slices of tissue obtained from the organ of interest (e.g., brain) are mounted on glass slides and incubated for a given time with a buffered radioligand solution. These sections are exposed to radiation‐sensitive film. Autoradiography may provide information about a radioligand’s suitability for PET studies with special regard to its affinity, selectivity, and nonspecific binding. An advantage compared with in vitro homogenate binding assays is the use of intact tissue, which provides information in an anatomical context. However, no data regarding the in vivo pharmacokinetics of a novel ligand can be deduced from such an experiment. This lack of information can be compensated for by ex vivo autoradiography approaches, where, as mentioned earlier, the analysis is done in vitro after in vivo administration of the radioligand into small animals. Traditionally, autoradiography experiments are performed with 3H‐ or 125 I‐labeled compounds, but the use of b‐sensitive film makes the procedure also suitable for molecules labeled with positron emitters, even though this lowers the achievable spatial resolution. An useful variant

. Table 3-3 Important receptor-mapping parameters in in vivo imaging of neurotrasmitter systems with PET Parameter In vitro target affinity Selectivity (relative affinity to competing binding sites) Concentration in binding sites Binding affinity Reversibility of binding Dissociation rate constant Equilibrium dissociation constant Target affinity/nonspecific affinity ratio Potency Toxicity Specific binding Nonspecific binding Plasma protein binding Blood–brain barrier permeability Equilibrium Tissue clearance Distribution volume Binding potential Occupancy Receptor saturation

Kd/Ki/IC50/Km

Unit nM

Bmax

pM/ml

k–1/koff/k4 koff Kd

sec–1, min–1

EC50 LD50

mM mg/kg Bmax

log P log P

BP %

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of this method is whole‐hemisphere autoradiography of the postmortem human brain. A comparison between this technique and PET imaging is shown in > Table 3-4.

. Table 3-4 A comparison of postmortem autoradiography (PMAR) and PET PMAR

PET

Advantage High spatial resolution Quantifiable Relatively easy pharmacology Easy receptor discrimination Behavior, disease versus distribution, binding

Disadvantage Postmortem Ante finem effects? (Drug treatment) Postmortem changes/deterioration? Static (snapshot) Limited availability Lower resolution In vivo limiting factors (blood–brain barrier, in vivo metabolism, etc.)

Quantifiable Test–retest possible Kinetic follow‐up Larger populations possible

Ligand‐dependent selectivity

The next step in receptor radioligand development is normally PET investigations in animals. For brain‐ imaging studies, nonhuman primates—such as cynomolgus monkey (Macaca fascicularis)—are preferred. Analysis of labeled metabolites from venous blood samples provides useful information regarding clearance and metabolic pathways. Administration of potent and selective competing compounds before radioligand injection (pretreatment) or during the time course of the PET experiment (displacement) can demonstrate the specificity and reversibility of radioligand binding (see later). It should be noted, however, that species differences may be encountered and lead to different results between animals and human subjects.

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Measuring Radioligand Effects in the Brain

The great advantage of the PET technique is that it is capable of obtaining absolute measurements of regional radioactivity concentrations which, in turn, with the help of appropriate kinetic models, can be transformed into quantitative parametric maps of related receptor parameters. What is relevant from our point of view is that several parameters related to the distribution and density of receptor systems and the ligands’ interaction with the receptors can also be measured (> Table 3-5). The initial selection of radioligands for neurotransmitter binding sites, such as receptors, neuronal uptake systems, and vesicular uptake systems, is often guided by data obtained in vitro by using tritiated or iodinated radioligands or by displacing a reference radioligand with the unlabeled molecule. In vitro binding normally provides information regarding ligand ‘‘affinity’’ (e.g., the dissociation equilibrium constants Kd or Ki) and ‘‘selectivity’’ (i.e., the relative affinity to competing binding sites) as well as regarding the ‘‘concentration’’ of binding sites (Bmax). The optimum affinity is closely related to the expected Bmax. It is preferable if the Bmax clearly exceeds the Kd of a ligand, i.e., if a binding site exists in vivo at nanomolar concentrations, a potentially successful radioligand ideally should have a subnanomolar affinity. Binding affinity is an important factor that determines the ratio of specific binding to nonspecific binding. The higher the ratio the more sensitive the signal is likely to be to changes in available binding site concentration, caused by disease or drug occupancy. Binding affinity (i.e., the fraction of dissociation rate constant, koff, and association rate constant, kon) usually governs the approach to be taken in the biomathematical modeling of the ligand–receptor

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. Table 3-5 Optimal criteria of PET ligands for neuroreceptors Aspect Radiochemical development

Biochemical properties

Favorable brain exposure

Key criteria  Should be radiolabeled with 11C or 18F  Radiolabeling should yield high specific radioactivity  Should be radiolabeled in a position not subject to formation of radioactive metabolites that pass the BBB  Sufficient affinity to the target receptor system (0.01–5 nM; the ideal affinity is dependent on target expression level)  High selectivity for the target receptor system (ideally 100‐fold as compared with any other binding sites)  Low nonspecific binding  As few as possible labeled metabolites in the brain; if they occur, their concentrations should be low with rapid clearance  Good permeation through the blood–brain barrier  Lipophilic (log P = 1–4)  Not a substrate for P‐glycoproteins  Low volume of distribution (Vd)  Low protein binding in plasma  No trapping in peripheral compartments

interaction. If the binding of a radioligand is reversible over the timescale of a PET experiment, ‘‘equilibrium’’ approaches toward quantification can be utilized. On the contrary, irreversible ligands normally demand ‘‘kinetic’’ modeling, wherein the transfer of radioligand between pharmacological compartments (e.g., plasma, tissue, receptors) is described in terms of rate constants. This approach requires in most cases the determination of an input function (i.e., the time course of free radioligand in plasma), which makes the measurement of radioligand metabolites in arterial plasma necessary. Very high binding affinity of a radioligand in combination with a comparatively slow clearance from tissue can restrict its usefulness for PET, as the rate‐limiting step of tracer retention may become the delivery instead of the binding process (flow‐limited conditions). A further important criterion for a radioligand is binding selectivity. Ideally, the affinity of a radioligand should be highest for the site of interest by more than one order of magnitude. However, lack of selectivity may be acceptable if nontarget sites are separated anatomically from the target‐binding sites. In the light of advances in molecular biology and pharmacology the term selectivity often needs to be revised. Most neurotransmitter receptors have now been found to exhibit multiple subtypes, and ligands that were initially thought to bind to a single class of receptors, truly display affinity toward several subtypes. This fact is also reflected by different ‘‘research philosophies’’ in drug development: whether ‘‘pharmacologically clean’’ molecules need to be developed with extremely high affinity to a given receptor system and low to other systems, therefore exhibiting high selectivity; or molecules with ‘‘rich pharmacology’’ are preferred, with an ‘‘optimal’’ mixture of affinities to a number of different receptor systems. Another substantial consideration in the development of a new radioligand is estimation of nonspecific binding. This is an essentially nonsaturable component of the total tissue uptake of a radioligand, usually attributed to adhesion to proteins and lipids. Nonspecific binding and its clearance in vivo are difficult to predict with absolute confidence. However, within a class of structurally related compounds, nonspecific interactions with tissue generally increase with increased lipophilicity. The logarithm of the partition coefficient between water (or preferably buffer to account for ionization at physiological pH) and octanol (log P) is often taken as a useful index for the lipophilicity of a compound in the context of biological systems. Conversely, some degree of lipid solubility is needed for good passage over the blood–brain barrier, which is a prerequisite for satisfactory counting statistics. However, the lipophilic nature of a molecule might also favor binding to plasma proteins, thus reducing the available ‘‘free fraction’’ in blood that is

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capable of diffusing through membranes. Moreover, lipophilic molecules can be extracted and metabolized in lung tissue when passing through the lung circulation, which prevents them from reaching their sites of action. Taken together, it appears that there is an optimal—but rather narrow—‘‘window’’ of lipophilicity for brain radioligands, wherein brain uptake is high and nonspecific binding comparatively weak. PET can quantitatively measure the regional radioactivity concentration without being able to distinguish the chemical forms or environments in which the radioactivity resides. For a clearly interpretable signal, it is therefore necessary that radiolabeled metabolites do not contribute to specific binding. Thus, radioligands should be preferably resistant to rapid metabolism over the period of data acquisition. Furthermore, radiolabeled metabolites should not be taken up and/or retained in the target area. This requirement may have important consequences concerning the elaboration of a radiolabeling strategy. A very important consideration in the context of radiochemistry is specific radioactivity (SR) of the radioligand. Too low SR may result in pharmacological effects or toxicity of the radiotracer. Moreover, low SR may saturate the biological system of interest, thus abolishing the mandatory tracer conditions. For low‐density binding sites very high SR is essential in order to exclude a substantial occupation of target sites by unlabeled ligand. The controlled administration of high SR radioligands versus low SR ligands may help estimate the binding profile of the labeled molecule as well as the general nature or actual status of the binding site. A major consideration is related to the intrinsic activity and efficacy of the prospective ligand as well as the developed radioligand on a given receptor system. The ligands have an intrinsic activity which is (1) 0 for antagonists fully blocking a receptor system; (2) vary between 0 and 1 for agonists (partial or full agonists); and (3) vary between 0 and –1 for inverse agonists (partial or full inverse agonists). The intrinsic efficacy of a ligand on a receptor can vary between a maximal effect and a minimal or no effect. An example for these properties is given in > Figure 3-7.

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Modeling Ligand Effects in the Brain

The great advantage of the PET technique, in contrast to other functional imaging modalities, is that it is capable of obtaining absolute measurements of regional radioactivity concentrations which, in turn, with the help of appropriate tracer kinetic models can be transformed into quantitative parametric maps of various biochemical, physiological, or neurotransmitter/receptor system‐related parameters. The quantitative measurements with PET require the determination of tissue and blood/plasma radioactivity concentrations and the a priori knowledge of a number of experimental parameters, including basic features related to the tracer, the biochemical and physiological characteristics, metabolic stability of the tracer/ligand, and those of the scanner. In the next step, a multicompartmental model describing the distribution and metabolism of the ligand in the brain and, eventually, in other body compartments, is developed, tested, and validated. With the help of appropriate tracer kinetic models the requested biological variables, e.g., receptor occupancy data, can be described in quantitative terms in precise anatomical context in the organ covered by the PET scan (> Figure 3-8).

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Two Approaches

The PET technique is sensitive for determinations of concentrations as low as the sub‐picomolar range (10–12 mol/l). Effective radiochemical labeling provides a radioligand with high SR, i.e., with a high ratio of radiolabeled to unlabeled drug molecules. A consequence is that i.v. injection of less than a microgram (mg) of the radiolabeled drug is sufficient for a PET study in man. The concept ‘‘tracer dose’’ is often used to emphasize the low mass, which does not induce drug effects. Similarly to neuropsychopharmacological drug development studies with PET, there are two possible experimental designs usable in mapping neuroreceptor systems. 1. The direct approach: to radiolabel the prospective ligand and measure its uptake, anatomical distribution, and binding in the brain. 2. The indirect approach: to study how an unlabeled molecule inhibits the specific binding of a well‐ characterized selective PET radioligand.

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. Figure 3-7 Intrinsic activity and effect of compounds labeled as existing or prospective PET radioligands for GABAA– benzodiazepine receptors. The ligands have an intrinsic activity which may change for agonists between 0 and 1, for inverse agonists between 0 and 1, and 0 for antagonists that can fully block a receptor. The intrinsic efficacy of a ligand on a receptor can also vary between maximal effect and no effect. In the case of benzodiazepine receptors, those ligands have positive intrinsic efficacy that increase GABAergic neurotransmission. The lower part of the figure shows recently developed compounds, a part of which are already used as antipsychotic drugs, whereas others are used as radioligands

7.1 Direct Approach: Radiolabeling of Ligands In this case the ligand can be radiolabeled and administered intravenously. The ligand’s brain uptake and distribution can be visualized and, alongside with a number of binding parameters, can be measured in a quantitative manner with PET (> Figure 3-9).

7.2 Indirect Approach: Using Radiolabeled Ligands and Drug Candidates In this case the unlabeled drug (ligand, drug, or drug candidate molecule) is competing with a well‐ characterized radioligand for occupying a receptor system (> Figure 3-10). The competition between the two molecules is not really a ‘‘fair competition’’, as the amount of the unlabeled drug exceeds that of the labeled drug. The unlabeled drug is given in pharmacological dose (mg range) whereas the labeled ligand is given in tracer dose (pico‐nano‐microgram range), the two doses being different in several orders of

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. Figure 3-8 The principles of quantitative PET imaging. The datasets required for acquiring quantitative data needed for modeling are: tissue radioactivity measurements, obtained with a PET scanner, and blood (and, eventually, plasma) radioactivity measurements, obtained by way of regular blood sampling. Quantitative models are usually compartmental models with the help of which biochemical parameters can be estimated in a qualitative manner

magnitude. This is done in order to occupy a given receptor system as completely as possible by the unlabeled drug and test the occupancy or ‘‘blockade’’ of the system. This can be achieved either by giving the unlabeled drug before the administration of the labeled drug (pretreatment) (see > Figure 3-11a) or after its administration (displacement) (> Figure 3-11b).

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Radioligands for Mapping Neurotransmitter Systems: Some Examples

A short overview of the central neuroreceptor systems with useful radioligands is given in > Table 3-6 and PET images, obtained in humans, of a few representative PET radioligands are shown in > Figure 3-12. Despite the fact that the monoamine neurotransmitter systems represent only a fraction of the central neuroreceptors, the most useful and best‐characterized PET radioligands are available for these systems; more specifically: the dopamine and serotonin systems. Intense studies have been focused in recent years to target other neuroreceptor systems, as well. Furthermore, PET radioligand development has also been focusing nowadays on the development of useful radiolabeled biomarkers for other than receptor binding sites in the brain, including amyloid plaques.

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. Figure 3-9 The direct approach involves the radiolabeling of a potential novel ligand and to trace its anatomical distribution and measure its binding in the brain

. Figure 3-10 A basic principle used in PET radioligand development. A well‐characterized labeled radioligand (used in tracer dose) is competing with a drug (used in therapeutic dose) for the same receptor system

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. Figure 3-11 The principles of pretreatment and displacement. The baseline curve (indicated with rhomboids) displays the normal kinetic behavior of a radioligand in the brain or a brain structure. If a drug, having affinity for the same receptor system and thereby blocking it, is administered before the radioligand injection in pharmacological dose, the radiolabeled ligand cannot bind to the blocked receptors in the same amount as in the baseline condition. This is called pretreatment condition (indicated with squares). If a drug, having affinity for the same receptor system and thereby blocking it, is administered after the radioligand injection in pharmacological dose (injection time is indicated by the arrow), the radiolabeled ligand, already bound to the receptors, will compete with the unlabeled drug. Because of the concentration differences between the labeled ligand and the unlabeled drug (tracer dose versus pharmacological dose; concentration differences usually over four orders of magnitude), the unlabeled drug will displace the radioligands bound to the receptors. This is called displacement condition (indicated with triangles)

8.1 Dopamine Receptor and Transporter Ligands Dopamine exerts its signaling effect by binding to specific membrane bound receptors, which belong to the family of G protein‐coupled receptors. Detailed molecular genetic studies have shown that the five types of dopamine receptors belong to two receptor families: D2, D3, and D4 receptors form one family, whereas D1 and D5 receptors form the other (> Table 3-7). Until now, only D1 and D2 receptors have been successfully visualized in vivo in humans, whereas there exist no useful selective radioligands for the dopamine D3 and D4 subtypes. There is a need to develop highly selective dopamine receptor radioligands for all the five subtypes, with special regard to the D3, D4, and D5 subtypes. The available dopamine radioligands labeled with b‐emitters have been extensively used in investigation of the dopamine receptors in physiology, neurological and mental disorders, and clinical pharmacology. Despite the fact that of the several billions of neurons in the human brain only a very small fraction (approximately 0.0003–0.0004%) use dopamine as a neurotransmitter; dopamine signaling plays a cardinal role in several brain functions, including locomotor control, positive reinforcement, cognitive functions, personality traits, and neuroendocrine regulation. Alterations of dopaminergic neurotransmission have been implicated in the pathophysiology of several neuropsychiatric disorders, such as Parkinson’s disease, Huntington’s disease, schizophrenia, ADHD, and drug abuse. The recently available highly useful PET radioligands of the dopamine receptor system use a few leads: the benzazepines such as [11C]SCH 23390 and [11C]NNC 112 bind to both D1 and D5 and the benzamides such as [11C]raclopride bind to both D2 and D3.

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. Table 3-6 Some representative PET radioligands used to study neuroreceptors in man Neurotransmitter systems Dopamine D1 Dopamine D2

Dopamine D3 Dopamine transporter

Serotonin 5‐HT1A

Serotonin 5‐HT2A

Serotonin transporter

Norepinephrine transporter

Glutamate Opiate

Muscarinic

Nicotinic

Radioligand [11C]‐SCH 23390 [11C]‐NNC 112 [11C]‐Raclopride [11C]‐NMSP [11C]‐FLB 457 [11C]‐MNPA [18F]‐Fallypride [18F]‐Fluoroethylspiperone [11C]‐RGH1756 [11C]‐MMC [11C]‐Methylphenidate [11C]‐PE2I [11C]‐b‐CIT‐FE [11C]‐b‐CFT [11C]‐Altropane [11C]‐Cocaine [18F]‐b‐CIT‐FP [Carbonyl‐11C]‐WAY‐100635 [11C]‐DWAY [11C]‐FCWAY [11C]‐NAD299 [11C]‐NMSP [11C]‐MDL 100907 [18F]‐Altanserin [18F]‐Setoperone [11C]‐DASB [11C]‐MADAM [11C]‐McN5652 [11C]‐nor‐b‐CIT [11C]‐MeNER

Selected references Halldin et al. (1986); Farde et al. (1987) Halldin et al. (1998a); Abi‐Dargham et al. (2000) Farde et al. (1986); Halldin et al. (1991) Wagner et al. (1983); Burns et al. (1984) Halldin et al. (1995); Olsson et al. (1999) Finnema et al. (2005); Seneca et al. (2005) Mukherjee et al. (1999); Christian et al. (2000) Jovkar et al. (1990); Wienhard et al. (1990) So´va´go´ et al. (2004, 2005) Gao et al. (2005) Ding et al. (1994); Volkow et al. (1998) Halldin et al. (1998a); Dolle´ et al. (2000) Halldin et al. (1996); Farde et al. (2000) Laakso et al. (1998a); Tsukada et al. (2001a) Madras et al. (1998); Fischman et al. (2001) Fowler et al. (1989); Volkow et al. (1999) Chaly et al. (1996); Lundkvist et al. (1997) Pike et al. (1996); Farde et al. (1998) Pike et al. (1998); Marchais‐Oberwinkler et al. (2005) Lang et al. (1999); Carson et al. (2000, 2002) Sandell et al. (1999, 2002); Andree et al. (2003) Burns et al. (1984); Andree et al. (1998) Lundkvist et al. (1996); Ito et al. (1998) Crouzel et al. (1992); Biver et al. (1994) Blin et al. (1990); Crouzel et al. (1992) Houle et al. (2000); Wilson et al. (2002) Halldin et al. (2005); Lundberg et al. (2005) Szabo´ et al. (1995, 1996) Mu¨ller et al. (1993); Bergstro¨m et al. (1997) Schou et al. (2003)

[18F]‐FMeNER [18F]‐FD2MeNER [11C]‐Desipramine [11C]‐Talopram [11C]‐Talsupram [11C]‐MPEP [11C]‐JNJ‐16567083) [11C]‐Diprenorphine [11C]‐Carfentanil [18F]‐Cyclofoxy [11C]‐NMPD [11C]‐3‐MPB [11C]‐Benztropine [11C]‐Nicotine [11C]‐Mecamylamine

Schou et al. (2004) Schou et al. (2005); Seneca et al. (2005) Van Dort et al. (1997); Schou et al. (2006) McConathy et al. (2004); Schou et al. (2006) Schou et al. (2004); McConathy et al. (2004) Yu et al. (2005) Huang et al. (2005) Mayberg et al. (1991); Jones et al. (1994) Dannals et al. (1985); Frost et al. (1989) Theodore et al. (1992); Cohen et al. (1998) Mulholland et al. (1995); Zubieta et al. (2001) Takahashi et al. (1999); Tsukada et al. (2001a) Dewey et al. (1990); Fujiwara et al. (1996) Nordberg et al. (1989) Sobrio et al. (2005)

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. Table 3-6 (continued) Neurotransmitter systems Histamine Adenosine Neurokinin‐1 GABA/benzodiazepine Peripheral benzodiazepine

Radioligand 2‐[18F]Fluoro‐A‐85380 [11C]‐Doxepin [11C]‐Pyrilamine [11C]‐MDPX [18F]‐CPFPX [18C]‐SPA‐RQ [11C]‐Flumazenil [11C]‐RO 5‐4513 [11C]‐PK11195 [11C]‐Vinpocetine [11C]‐DAA1106

Selected references Gallezot et al. (2004); Obrzut et al. (2005) Tashiro et al. (2004); Iwabuchi et al. (2005) Szabo´ et al. (1993); Kim et al. (1999) Fukumitsu et al. (2003, 2005); Kimura et al. (2004) Bauer et al. (2003); Meyer et al. (2006) Bergstro¨m et al. (2004); Solin et al. (2004); Hietala et al. (2005) Mazie`re et al. (1984); Persson et al. (1989) Halldin et al. (1992); Pike et al. (1993) Hashimoto et al. (1989); Banati et al. (1999) Gulya´s et al. (2005); Vas and Gulya´s, (2005) Okuyama et al. (1999); Maeda et al. (2004)

A benzazepine derivative, SCH 23390, has been described as the first high‐affinity selective dopamine D1 receptor antagonist and labeled with positron‐emitting 11C. An improved benzazepine compound [11C]NNC 112 has also been developed and is now used worldwide. The gold standard of dopamine D2 receptor ligands is 11C‐raclopride, which has permitted the selective analysis by PET of central D2 receptors in primates and humans. Radiolabeled benzamides of picomolar affinities, which make them suitable to examine the extrastriatal D2 receptors in the human brain, have been developed during the recent years, such as [11C]FLB 457. The butyrophenon N‐[11C]methylspiperone ([11C]NMSP) is also showing promising features. Whereas these ligands are antagonists, the newly developed agonist radioligand, [11C]MNPA, shows high binding affinity to the D2 receptor system. An 18F‐labeled benzamide, also suitable for investigation of extrastriatal D2 receptors, is 18F‐fallypride. The labeling of dopamine D3 receptors is a challenge, and whereas a few candidates have been tested (e.g., [11C]‐RGH1756), no selective radioligand is available at this time. The dopamine reuptake sites play an effective role in the regulation of the intrasynaptic dopamine concentration. Several attempts have been made to find appropriate PET ligands for the labeling of this system. The recently used ligands include [11C]‐PE2I (N‐(3‐iodoprop‐2E‐enyl)‐2b‐carbomethoxy‐3b‐(40 ‐ methylphenyl)nortropane) and [11C]‐FE‐CIT (N‐(2‐fluoroethyl)‐2 b‐carbomethoxy‐3b‐(4‐iodophenyl) nortropane). An overview of the useful dopamine receptor and transporter radioligands are shown in > Figure 3‐12a.

8.2 Serotoninergic Neurotransmission Radioligands In addition to the dopamine system, the serotonin (5‐hydroxytryptamine, 5‐HT) system is the other monoaminergic system for which a number of PET radioligands are available. The serotonin system consists of a large number of postsynaptic 5‐HT receptors, belonging to seven major families (5‐HT1–5‐HT7), each containing several subtypes, and the serotonin transporter, regulating the reuptake of the intrasynaptic serotonin into the presynaptic neuron. Within the 5HT1 group there are subtypes 5HT1A, 5HT1B, 5HT1D, 5HT1E, and 5HT1F. There are three 5HT2 subtypes, 5HT2A, 5HT2B, and 5HT2C as well as two 5HT5 subtypes, 5HT5B and 5HT5B. Most of these receptors are metabotropic; however, the 5HT3 class is, for instance, ionotropic. The most commonly studied serotonin receptor subsystem, the 5‐HT1A, is present in high densities in the hippocampus, septum, amygdala, hypothalamus, and neocortex. The other commonly studied receptor subsystem, the 5‐HT2A receptors, are present in the neocortex, followed by, in the hippocampus, basal ganglia, and thalamus. The transporter has high densities in midbrain structures, with special regard to the raphe nuclei, the thalamus, and the striatum.

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. Figure 3-12 Horizontal PET images of the human brain showing the cerebral uptake of some of the most commonly used PET radioligands. For details. see text and > Table 3-6

Radioligand development to all receptor subtypes is a challenge and a number of PET laboratories are working on this challenge. The recently available radioligands include the 5‐HT1A antagonist WAY‐100635 (N‐(2‐(4‐(2‐methoxyphenyl)‐1‐piperazinyl)ethyl)‐N‐(2‐pyridyl) cyclohexane‐carboxamide), the 5‐HT2A antagonists MDL100907 ((R)‐(þ)‐a‐(2,3‐dimethoxyphenyl)‐1‐[2‐(4‐fluoro‐phenyl)ethyl]‐4‐piperidine methanol] and N‐methylspiperone (NMSP), and the transporter ligands MADAM (11C‐N,N‐Dimethyl‐2‐ (2‐amino‐4‐methylphenylthio)benzylamine) and DASP (3‐amino‐4‐(2‐dimethylaminomethyl‐phenylsulfanyl)benzonitrile) (see > Figure 3‐12b).

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. Table 3-7 Human dopamine receptor subtypes Amino acid

D1 446

Chromosome Introns G‐protein Coupling

5q31–34 No Gs þcAMP þIP3

Main localization in the CNS

Nucleus Caudatus Putamen

D2 D2A: 443 D2B: 414 11q22–q23 Yes Gi, Go cAMP Ca2þ channel þKþ channel IP3? Caudatus Putamen

Accumbens

Accumbens

Substantia nigra Cortex

Substantia nigra

D3 400

D4 387

D5 477

3q13.3 Yes Gi? cAMP?

11p15.5 Yes Gi cAMP?

4p15.1–16.1 No Gs þcAMP

Island of Calleja Accumbens

Entorhinal cortex

Hippocampus

Bed nucleus Stria terminalis

Hippocampus

Lateral mamillary bodies

Amygdala Medulla

8.3 Norepinephrine Neurotransmission Radioligands The third major family of CNS monoaminergic receptors is the noradrenaline or norepinephrine system. The radiochemistry of norepinephrine transporter ligands is, at this day, more developed that that of the receptor ligands. Among others, [11C]]MRB ([11C]O‐methylreboxetine), [11C]desipramine, and the 11C and 18F versions of MeNER ((S,S)‐2‐(a‐(2‐fluoromethoxyphenoxy)benzyl)morpholine), an O‐methyl analog of reboxetine, have recently been introduced to the PET community (see > Figure 3-12c).

8.4 Acetylcholine Radioligands The acetylcholine receptors are usually subdivided into nicotinic and muscarinic receptors, the former belonging to the ligand‐gated ion channels, whereas the latter being metabotropic receptors. Both systems have been implicated in several neurological functions and, consequently, in neurological or psychiatric diseases, including Alzheimer’s disease, Parkinson’s disease, and schizophrenia. Intensive radioligand development activities aim at targeting both systems. Despite this fact, no highly useful and widely used PET ligands exist for these two systems, despite the fact that a number of radioligands have recently been advocated, including [11C]NMPB, [11C]benztropin, [11C]scopolamine, [11C]IQNP, and [11C]IQNB for the muscarinic and [18F]2‐F‐A85380 and [11C]A84543 for the nicotinic systems.

8.5 Central Benzodiazepine‐Binding Site Ligands In the mammalian brain, two different benzodiazepine‐binding sites, the central and the peripheral types, have been characterized. The central type benzodiazepine‐binding site is part of the GABA/benzodiazepine/ Cl supramolecular receptor complex. This specific site mediates all pharmacological properties of

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the classical benzodiazepines (sedative, anxiolytic, anticonvulsant, and myorelaxant). The chloride channel‐ gating function of GABAA receptors in the CNS can be allosterically modulated by ligands acting at the benzodiazepine receptor. The compounds which have been labeled for visualizing central benzodiazepine receptors belong to three different structural chemical families: the classical benzodiazepines, the cyclopyrrolones, and the imidazobenzodiazepine derivatives. Several benzodiazepine agonists (flunitrazepam, diazepam, suriclone) have been prepared and used as radioligands. [11C]‐Flunitrazepam and [11C]‐diazepam led to the first visualization of benzodiazepine receptors in baboon and human brains. [11C]‐Suriclone, a cyclopyrrolone, has also been proposed for in vivo purposes. A search for a more specific central type benzodiazepine ligands has led to the labeling of antagonists such as oxoquazepam, a selective BZ1 subtype ligand, and flumazenil (RO151788), an imidazobenzodiazepine. Flumazenil has been labeled with 11C either by N‐methylation with [11C]‐methyl iodide or by esterification with [11C]‐ethyl iodide. An analogue of flumazenil labeled with 18F, [18F]‐fluoroethylflumazenil, appears to be a suitable PET ligand despite its lower affinity and more rapid metabolism and kinetics. [11C]‐Flumazenil is considered now as the reference tracer used for pharmacological and clinical PET studies of the central benzodiazepine receptor.

8.6 Peripheral Benzodiazepine Receptor or Benzodiazepine‐Binding Site Receptor Ligands Unlike the central benzodiazepine receptors which are located on the cell membrane, the peripheral benzodiazepine receptor (PBR) is localized in the mitochondrial and nuclear subcellular fractions. For this and other characteristics of the protein complex, it is different from the classical membrane receptors and is often termed as the peripheral benzodiazepine‐binding site (PBBS). The peripheral benzodiazepine receptor plays a cardinal role in cellular respiration, oxidative metabolism and ion transport, neurosteroid biosynthesis, porphyrin transport and heme synthesis, regulation of calcium flow, apoptosis, and several other cellular processes. In the brain, the PBR can be visualized in activated microglia and astrocytes. The isoquinoline PK11195, a ligand for the PBBS, binds with relative cellular selectivity to activated, but not resting, microglia. [11C]‐PK11195 has then be used to study inflammatory and neurodegenerative brain diseases in vivo using PET. Moreover, [11C]‐PK11195 appears to be a biomarker of neuronal injury not only at the primary lesion site but also at the antero‐ and retrograde projection areas of the lesioned neurons. More recently, other candidate radiomarkers have also been tested, including [11C]‐DAA1106 (N‐5‐fluoro‐2‐phenoxyphenyl)‐N‐(2‐hydroxy‐5‐methoxybenzyl)acetamide and [11C]‐vinpocetine (cis‐ ethyl‐apovincaminate). Vinpocetine enters the brain in larger proportions that PK11195, and with the help of it even the age‐related physiological increase of PBR densities in the human brain can be demonstrated (> Figure 3-12d).

8.7 Glutamate Neurotransmission Radioligands Metabotropic glutamate (mGlu) receptors play a vital role in normal brain functions, and consequently, also in neurological and psychiatric disorders. The precise functions of these receptors are still undefined. Progress toward understanding their functions has been hampered by the lack of selective ligands with appropriate pharmacokinetic properties. However, the glutamate system is a conundrum for PET radiochemistry. Despite years of intensive search for useful radioligands, no breakthrough has yet been reported in this field. A few promising approaches were published, using, among others, 2‐methyl‐6‐(phenylethynyl)‐ pyridine (MPEP) and its analogues, M‐MPEP and M‐PEPy, [11C]‐ABP688 (3‐(6‐methyl‐pyridin‐2‐ylethynyl)‐cyclohex‐2‐enone‐O‐[11C]‐methyl‐oxime), bis(phenylalkyl)amines, [11C]‐3‐[2‐[(3‐methoxyphenylamino)‐carbonyl]‐ethenyl]‐4,6‐dichloroindole‐2‐carboxylic acid (3MPICA), and N,N0 ‐diphenyl and

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3

N‐naphthyl‐N0 ‐phenyl guanidine derivatives. As none of these approaches have until now resulted in a usable PET radiomarker, this particular field is one of the greatest challenges nowadays of PET radioligand chemistry.

8.8 Cannabinoid Neurotransmission Radioligands In the endocannabinoid system two receptor subtypes, CB1 and CB2, are recognized. Both receptors belong to the G protein‐coupled superclass of receptors. CB1 receptors are located throughout the body including within the CNS at presynaptic nerve terminals. CB2 receptors are mainly associated with cells of the immune system. Brain CB1 receptors represent an interesting target for the treatment of several psychiatric (e.g., anxiety, addiction, depression) and neurodegenerative disorders (e.g., Huntington’s disease and Tourette’s syndrome). The role of the CB1 receptors in these disorders is not well understood. The search for adequate PET radioligands is recently centered around a few structures, including 1,5‐diarylpyrazole, 3‐(4‐fluoronaphthoyl)‐1‐(N‐methylpiperidin‐2‐ylmethyl)indole, N‐([18F]fluorophenyl)‐ 5‐(4‐bromophenyl)‐1‐(2,4‐dichlorophenyl)‐1H‐pyrazole‐3‐carboxamide, and O‐methyl‐[11C]‐1‐(2‐chlorophenyl)‐5‐(4‐methoxyphenyl)‐4‐methyl‐1H‐pyrazole‐3‐carboxylic acid piperidin‐1‐ylamide ([11C]‐1).

8.9 Opioid Receptors The opioid receptors belong to the G protein‐coupled receptor families. The opioid receptors, m, d and k, are the mediators of the pharmacological effects of opioid drugs. The receptor subtypes are broadly expressed in the CNS. Their activation by opioid compounds (including morphine, codeine, heroin) is intimately involved with reward, tolerance, and withdrawal. Their significant role in analgesia, antinociception, and drug addiction has been demonstrated extensively. Among the recently available PET radioligands, [11C]‐carfentanil, a m‐receptor agonist, has been studied most extensively. Furthermore, [11C]‐diprenorphine, a nonspecific antagonist, [18F]‐diprenorphine, a nonspecific antagonist, and [18F]‐cyclofoxy, a m‐ and k‐receptor antagonist, are worth mentioning. Ongoing research focuses on other structures, as well, including cyclohexyl piperazine, diprenorphine, benzamide analogues, and 4‐anilidopiperidines.

9

Conclusion

PET radioligands play an important role in the qualitative visualization and quantitative exploration and mapping of neuroreceptor systems in the primate brain. With the help of PET imaging using adequate radioligands, the distribution of various receptor systems in the brain, their densities, and occupancy levels can be quantitatively measured in precise anatomical context. Physiological situations can be explored in normal conditions and under various pharmacological challenges, as well as alterations related to disease conditions or long‐lasting drug therapies or addiction can be studied. PET will present for the coming years the ‘‘par excellence’’ research tool for studies aiming at the in vivo quantitative exploration and anatomical mapping of neuroreceptor systems in the human brain.

References Abi‐Dargham A, Martinez D, Mawlawi O, Simpson N, Hwang DR, et al. 2000. Measurement of striatal and extrastriatal dopamine D1 receptor binding potential with [11C]NNC 112 in humans: Validation and reproducibility. J Cereb Blood Flow Metab 20(2): 225-243.

Andree B, Nyberg S, Ito H, Ginovart N, Brunner F, et al. 1998. Positron emission tomographic analysis of dose-dependent MDL 100,907 binding to 5-hydroxytryptamine-2A receptors in the human brain. J Clin Psychopharmacol 18: 317-323.

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Andree B, Hedman A, Thorberg SO, Nilsson D, Halldin C, et al. 2003. Positron emission tomographic analysis of dose‐ dependent NAD‐299 binding to 5‐hydroxytryptamine‐1A receptors in the human brain. Psychopharmacology (Berl) 167(1): 37-45. Banati RB, Goerres GW, Myers R, Gunn RN, Turkheimer FE, et al. 1999. [11C](R)‐PK11195 positron emission tomography imaging of activated microglia in vivo in Rasmussen’s encephalitis. Neurology 53(9): 2199-2203. Bauer A, Holschbach MH, Meyer PT, Boy C, Herzog H, et al. 2003. In vivo imaging of adenosine A1 receptors in the human brain with [18F]CPFPX and positron emission tomography. Neuroimage 19(4): 1760-1769. Bergstro¨m KA, Halldin C, Hall H, Lundkvist C, Ginovart N, et al. 1997. In vitro and in vivo characterisation of nor‐b‐ CIT: A potential radioligand for visualisation of the serotonin transporter in the brain. Eur J Nucl Med 24(6): 596-601. Bergstro¨m M, Hargreaves RJ, Burns HD, Goldberg MR, Sciberras D, et al. 2004. Human positron emission tomography studies of brain neurokinin 1 receptor occupancy by aprepitant. Biol Psychiatry 55(10): 1007-1012. Biver F, Goldman S, Luxen A, et al. 1994. Multicompartmental study of fluorine‐18 altanserin binding to brain 5HT2 receptors in humans using positron emission tomography. Eur J Nucl Med 21: 937-946. Blin J, Sette G, Fiorelli M, Bletry O, Elghozi JL, et al. 1990. A method for the in vivo investigation of the serotonergic 5‐HT2 receptors in the human cerebral cortex using positron emission tomography and 18F‐labeled setoperone. J Neurochem 54(5): 1744-1754. Burns HD, Dannals RF, Langstro¨m B, et al. 1984. (3‐N‐[11C]‐ methyl)spiperone—a ligand binding to dopamine receptors: Radiochemical synthesis and biodistribution studies in mice. J Nucl Med 25: 1222-1227. Burns HD, Dannals RF, Langstrom B, Ravert HT, Zemyan SE, et al. 1984. (3‐N‐[11C]methyl)spiperone, a ligand binding to dopamine receptors: Radiochemical synthesis and biodistribution studies in mice. J Nucl Med 25(11): 1222-1227. Carson RE, Toczek MT, Lang L, et al. 2002. Human functional imaging with the 5‐HT1A ligand 18F‐FCWAY. J Nucl Med 43: 55. Carson RE, Lang L, Watabe H, Der MG, Adams HR, et al. 2000. PET evaluation of [(18)F]FCWAY, an analog of the 5-HT(1A) receptor antagonist, WAY-100635. Nucl Med Biol 27: 493-497. Chaly T, Dhawan V, Kazumata K, et al. 1996. Radiosynthesis of [18F]N‐3‐fluoropropyl‐2‐b‐carbomethoxy‐3‐b‐(4iodophenyl)nortropane and the first human study with positron emission tomography. Nucl Med Biol 23: 999-1004. Christian BT, Shi B, Mukherjee J. 2000. Quantification of striatal and extrastriatal D2 dopamine receptors using

PET imaging of [18F]fallypride in nonhuman primates. Synapse 38: 71-79. Cohen RM, Carson RE, Channing M, et al. 1998. [18F]‐cyclofoxy PET imaging in man. J Nucl Med 29: 796-807. Crouzel C, Guillaume M, Barre L, Lemaire C, Pike VW. 1992. Ligands and tracers for PET studies of the 5‐HT system— Current status. Nucl Med Biol‐Int J Rad Appl B 19: 857-870. Dannals RF, Ravert HT, Frost JJ, Wilson AA, Burns HD, et al. 1985. Radiosynthesis of an opiate receptor binging radiotracer: (11C) Carfentanil. Int J Appl Radiat Isot 36: 303-306. Dewey SL, Mac Gregor RR, Brodie JD, et al. 1990. Mapping muscarinic receptors in human and baboon brain using [N‐11C‐methyl]‐benztropine. Synapse 5: 213-223. Ding YS, Fowler JS, Volkow ND, Gatley SJ, Logan J, et al. 1994. Pharmacokinetics and in vivo specificity of [11C]DL‐threo‐ methylphenidate for the presynaptic dopaminergic neuron. Synapse 18(2): 152-160. Dolle F, Bottlaender M, Demphel S, et al. 2000. High efficient synthesis of [11C]PE2I, a selective radioliogand for the quantification of the dopamine transporter using PET. J Label Compd Radiopharm 43: 997-1004. Farde L, Ginovart N, Halldin C, Chou YH, Olsson H, et al. 2000. A PET study of [11C]b‐CIT‐FE binding to the dopamine transporter in the monkey and human brain. Int J Neuropsychopharm 2: 203-214. Farde L, Hall H, Ehrin E, Sedvall G. 1986. Quantitative analysis of dopamine D2 receptor binding in the human brain by PET. Science 231: 258-261. Farde L, Halldin C, Stone‐Elander S, Sedvall G. 1987. PET analysis of human dopamine receptor subtypes using 11 C‐SCH 23390 and 11C‐raclopride. Psychopharmacology 92: 278-284. Farde L, Ito H, Swahn C‐G, Pike VW, Halldin C. 1998. Quantitative analyses of carbonyl‐carbon‐11‐WAY‐100635 binding to central 5‐hydroxytryptamine‐1A receptors in man. J Nucl Med 39: 1965-1971. Finnema SJ, Seneca N, Farde L, Shchukin E, Sovago J, et al. 2005. A preliminary PET evaluation of the new dopamine D2 receptor agonist [11C]MNPA in cynomolgus monkey. Nucl Med Biol 32(4): 353-360. Fischman AJ, Bonab AA, Babich JW, Livni E, Alpert NM, et al. 2001. [11C, 127I] Altropane: A highly selective ligand for PET imaging of dopamine transporter sites. Synapse 39(4): 332-342. Fowler JS, Volkow ND, Wang GJ, Ding YS, Dewey SL. 1999. PET and drug research and development. J Nucl Med 40(7): 1154-1163. Fowler JS, Volkow ND, Wolf AP, Dewey SL, Schlyer DJ, et al. 1989. Mapping cocaine binding sites in human and baboon brain in vivo. Synapse4(4): 371-377. Frost JJ, Douglass KH, Mayberg HS, et al. 1989. Multicompartmental analysis of [11C]‐carfentanil binding to opiate

In vivo imaging of neurotransmitter systems with PET receptors in humans measured by positron emission tomography. J Cereb Blood Flow Metab 9: 398-409. Fujiwara T, Meija M, Itoh M, Anai K, Meguro K, et al. 1996. Quantitative imaging of [11C]benztropine in the human brain with graphic analysis and spectral analysis. Quantification of Brain Function Using PET. Myers R, Cunningham MG, Bailey D, Jones T, editors. San Diego: Academic Press. Fukumitsu N, Ishii K, Kimura Y, Oda K, Sasaki T, et al. 2003. Imaging of adenosine A1 receptors in the human brain by positron emission tomography with [11C]MPDX. Ann Nucl Med 17(6): 511-515. Fukumitsu N, Ishii K, Kimura Y, Oda K, Sasaki T, et al. 2005. Adenosine A1 receptor mapping of the human brain by PET with 8‐dicyclopropylmethyl‐1‐11C‐methyl‐3‐propylxanthine. J Nucl Med 46(1): 32-37. Gallezot JD, Bottlaender M, Gregoire MC, Roumenov D, Deverre JR, et al. 2004. In vivo imaging of human cerebral nicotinic acetylcholine receptors with 2‐18F‐fluoro‐A‐85380 and PET. J Nucl Med 46(2): 240-247. Gao M, Mock BH, Hutchins GD, Zheng QH. 2005. Synthesis and initial PET imaging of new potential dopamine D3 receptor radioligands (E)‐4,3,2‐[11C]methoxy‐N‐4‐(4‐(2‐methoxyphenyl)piperazin‐1‐yl)butyl‐cinnamoylamides. Bioorg Med Chem 13(22): 6233-6243. Gulya´s B, Halldin C, Vas A, Banati RB, Shchukin E, et al. 2005. [11C]Vinpocetine: A prospective peripheral benzodiazepine receptor ligand for primate PET studies. J Neurol Sci 229–230: 219-223. Halldin C, Farde L, Hogberg T, et al. 1991. A comparative PET‐study of five carbon‐11 or fluorine‐18 labelled salicylamides. Preparation and in vitro dopamine D2 receptor binding. Nucl Med Biol 18: 871-881. Halldin C, Farde L, Hogberg T, et al. 1995. Carbon‐11‐FLB 457: A radioligand for extrastriatal D2 dopamine receptors. J Nucl Med 36: 1275-1281. Halldin C, Farde L, Litton JE, Hall H, Sedvall G. 1992. (C‐11) Ro 15‐4513, a ligand for visualization of benzodiazepine receptor binding—Preparation, autoradiography and positron emission tomography. Psychopharmacology 108: 16-22. Halldin C, Farde L, Lundkvist C, et al. 1996. (11C)‐b‐CIT‐FE, a radioligand for quantitation of the dopamine transporter in the living brain using positron emission tomography. Synapse 22: 386-390. Halldin C, Foged C, Chou YH, et al. 1998a. Carbon‐11 NNC 112: A radioligand for PET examination of striatal and neocortical D1‐dopamine receptors. J Nucl Med 39: 2064-2068. Halldin C, Guilloteau D, Okubo Y, 1998b. PET‐characterization of [C‐11]PE2I binding to dopamine transporters in the monkey brain. J Nucl Med 39: 118.

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Halldin C, Gulya´s B, Farde L. 2001. PET studies with carbon‐ 11 radioligands in neuropsychological drug development. Curr Radiopharm Design 7: 1907–1929. Halldin C, Gulya´s B, Farde L. 2004. PET for drug development. From Morphological Imaging to Molecular Targeting. Berlin Heidelberg: Springer Verlag; pp. 95–110. Halldin C, Gulya´s B, Langer O, Farde L. 2001. Brain radioligands—State of the art and new trends. Q J Nucl Med 2: 139-152. Halldin C, Ho¨gberg T. 1991. Radiotracers: Synthesis and use in imaging. A Textbook of Drug Design and Development. Kroogsgaard‐Larsen P, Liljefors T, Madsen U, editors. Amsterdam: Harwood; pp. 174–205. Halldin C, Lundberg J, Sovago J, Gulya´s B, Guilloteau D, et al. 2005. [11C]MADAM, a new serotonin transporter radioligand characterized in the monkey brain by PET. Synapse 58(3): 173-183. Halldin C, Stone‐Elander S, Farde L, et al. 1986. Preparation of 11C‐labelled SCH 23390 for the in vivo study of dopamine D1 receptors using positron emission tomography. Appl Radiat Isot‐Int J Rad A 37: 1039-1043. Hashimoto K, Inoue O, Suzuki K, Yamasaki T, Kojima M. 1989. Synthesis and evaluation of 11C‐PK 11195 for in vivo study of peripheral‐type benzodiazepine receptors using positron emission tomography. Ann Nucl Med 3(2): 63-71. Heiss WD, Herholz K. 2006. Brain receptor imaging. J Nucl Med 47(2): 302-312. Hietala J, Nyman MJ, Eskola O, Laakso A, Gronroos T, et al. 2005. Visualization and quantification of neurokinin‐1 (NK1) receptors in the human brain. Mol Imaging Biol 7(4): 262-272. Houle S, Ginovart N, Hussey D, Meyer JH, Wilson AA. 2000. Imaging the serotonin transporter with positron emission tomography: Initial human studies with [11C]DAPP and [11C]DASB. Eur J Nucl Med 27(11): 1719-1722. Huang Y, Narendran R, Bischoff F, Guo N, Zhu Z, et al. 2005. A positron emission tomography radioligand for the in vivo labeling of metabotropic glutamate 1 receptor: (3‐ethyl‐2‐[ 11C]methyl‐6‐quinolinyl)(cis‐4‐methoxycyclohexyl)methanone. J Med Chem 48(16): 5096-5099. Ito H, Nyberg S, Halldin C, Lundkvist C, Farde L. 1998. PET imaging of central 5‐HT2A receptors with carbon‐11‐MDL 100,907. J Nucl Med 39(1): 208-214. Iwabuchi K, Ito C, Tashiro M, Kato M, Kano M, et al. 2005. Histamine H1 receptors in schizophrenic patients measured by positron emission tomography. Eur Neuropsychopharmacol 15(2): 185-191. Jones AK, Cunningham VJ, Ha‐Kawa SK, Fujiwara T, Liyii Q, et al. 1994. Quantitation of [11C]diprenorphine cerebral kinetics in man acquired by PET using presaturation, pulse‐chase and tracer‐only protocols. J Neurosci Methods 51(2): 123-134.

97

98

3

In vivo imaging of neurotransmitter systems with PET

Jovkar S, Wienhard K, Pawlik G, Coenen HH. 1990. The quantitative analysis of D2‐dopamine receptors in baboon striatum in vivo with 3‐N‐[20 ‐18F]fluoroethylspiperone using positron emission tomography. J Cereb Blood Flow Metab 10(5): 720-726. Kim SE, Szabo´ Z, Seki C, Ravert HT, Scheffel U, et al. 1999. Effect of tracer metabolism on PET measurement of [11C] pyrilamine binding to histamine H1 receptors. Ann Nucl Med 13(2): 101-107. Kimura Y, Ishii K, Fukumitsu N, Oda K, Sasaki T, et al. 2004. Quantitative analysis of adenosine A1 receptors in human brain using positron emission tomography and [1‐methyl‐ 11C]8‐dicyclopropylmethyl‐1‐methyl‐3‐propylxanthine. Nucl Med Biol 31(8): 975-981. Laakso A, Bergman J, Haaparanta M, Vilkman H, Solin O, et al. 1998. [F‐18]CFT ([F‐18]WIN 35,428), a radioligand to study the dopamine transporter with PET: Characterization in human subjects. Synapse 28: 244-250. Lang L, Jagoda E, Schmall B, Vuong BK, Adams HR, et al. 1999. Development of fluorine‐18‐labeled 5‐HT1A antagonists. J Med Chem 42(9): 1576-1586. Lundberg J, Odano I, Olsson H, Halldin C, Farde L. 2005. Quantification of 11C‐MADAM binding to the serotonin transporter in the human brain. J Nucl Med 46(9): 1505-1515. Lundkvist C, Halldin C, Ginovart N, et al. 1996. [C‐11]MDL 100907, a radioligand for selective imaging of 5‐HT2A receptors with positron emission tomography. Life Sci 58: PL187-PL192. Lundkvist C, Halldin C, Ginovart N, Swahn C‐G, Farde L. 1997. [18F]b‐CIT‐FP is superior to [11C]b‐CIT‐FP for quantitation of the dopamine transporter. Nucl Med Biol 24: 621-627. Madras BK, Meltzer PC, Liang AY, Elmaleh DR, Babich J, et al. 1998. Altropane, a SPECT or PET imaging probe for dopamine neurons: I. Dopamine transporter binding in primate brain. Synapse 29: 93-104. Maeda J, Suhara T, Zhang MR, Okauchi T, Yasuno F, et al. 2004. Novel peripheral benzodiazepine receptor ligand [11C]DAA1106 for PET: An imaging tool for glial cells in the brain. Synapse 52(4): 283-291. Marchais‐Oberwinkler S, Nowicki B, Pike VW, Halldin C, Sandell J, et al. 2005. N‐oxide analogs of WAY‐100635: New high affinity 5‐HT1A receptor antagonists. Bioorg Med Chem 13(3): 883-893. Mayberg HS, Sadzot B, Meltzer CC, Fisher RS, Lesser RP, et al. 1991. Quantification of mu and non‐mu opiate receptors in temporal lobe epilepsy using positron emission tomography. Ann Neurol 30(1): 3-11. Mazie`re M, Hantraye P, Prenant C, Sastre J, Comar D. 1984. Synthesis of ethyl 8‐fluoro‐5,6‐dihydro‐5‐[11C]methyl‐6‐ oxo‐4H‐imidazo [1,5‐a] [1,4]benzodiazepine‐3‐carboxylate

(RO 15.1788‐11C): A specific radioligand for the in vivo study of central benzodiazepine receptors by positron emission tomography. Int J Appl Radiat Isot 35: 973-976. McConathy J, Owens MJ, Kilts CD, Malveaux EJ, Camp VM, et al. 2004. Synthesis and biological evaluation of [11C] talopram and [11C]talsupram: Candidate PET ligands for the norepinephrine transporter. Nucl Med Biol 31(6): 705-718. Meyer PT, Elmenhorst D, Matusch A, Winz O, Zilles K, et al. 2006. 18F‐CPFPX PET: On the generation of parametric images and the effect of scan duration. J Nucl Med 47(2): 200-207. Mukherjee J, Yang ZY, Brown T, Lew R, Wernick M, et al. 1999. Preliminary assessment of extrastriatal dopamine D-2 receptor binding in the rodent and nonhuman primate brains using the high affinity radioligand, 18F-fallypride. Nucl Med Biol 26: 519-527. Mulholland GK, Kilbourn MR, Sherman P, et al. 1995. Synthesis, in vivo biodistribution and dosimetry of [C‐11]N‐ methylpiperidyl benzilate ([C‐11]NMPB), a muscarinic acetylcholine receptor antagonist. Nucl Med Biol 22: 13-17. Mu¨ller L, Halldin C, Farde L, Karlsson P, Hall H, et al. 1993. [11C] b‐CIT, a cocaine analogue. Preparation, autoradiography and preliminary PET investigations. Nucl Med Biol 20(3): 249-255. Nordberg A, Hartvig P, Lundqvist H, Antoni G, Ulin J, et al. 1989. Uptake and regional distribution of (þ)‐(R)‐ and ()‐(S)‐N‐[methyl‐11C]‐nicotine in the brains of rhesus monkey. An attempt to study nicotinic receptors in vivo. J Neural Transm Park Dis Dement Sect 1(3): 195-205. Obrzut SL, Koren AO, Mandelkern MA, Brody AL, Hoh CK, et al. 2005. Whole‐body radiation dosimetry of 2‐[18F] Fluoro‐A‐85380 in human PET imaging studies. Nucl Med Biol 32(8): 869-874. Okuyama S, Chaki S, Yoshikawa R, Ogawa S, Suzuki Y, et al. 1999. Neuropharmacological profile of peripheral benzodiazepine receptor agonists, DAA1097 and DAA1106. Life Sci 64(16): 1455-1464. Olsson H, Halldin C, Swahn C‐G, Farde L. 1993. Quantification of [C‐11]FLB 457 binding to extrastriatal dopamine receptors in the human brain. J Cerebr Blood Flow Metab 19: 1164-1173. Persson A, Pauli S, Halldin C, et al. 1989. Saturation analysis of specific 11C‐Ro 15–1788 binding to the human neocortex using positron emission tomography. Hum Psychopharmacol 4: 21-31. Pike VW, Halldin C, Crouzel C, et al. 1993. Radioligands for PET studies of central benzodiazepine receptors and PK (peripheral benzodiazepine) binding sites—Current status. Nucl Med Biol 20: 503-525. Pike VW, Halldin C, McCarron JA, Lundkvist C, Hirani E, et al. 1998. [Carbonyl‐11C]Desmethyl‐WAY‐100635 (DWAY)

In vivo imaging of neurotransmitter systems with PET is a potent and selective radioligand for central 5‐HT1A receptors in vitro and in vivo. Eur J Nucl Med 25(4): 338-346. Pike VW, Mccarron JA, Lammertsma AA, et al. 1996. Exquisite delineation of 5‐HT1A receptors in human brain with PET and [carbonyl‐C‐11]WAY‐100635. Eur J Pharmacol 301: R5-R7. Sandell J, Halldin C, Chou YH, Swahn CG, Thorberg SO, et al. 2002. PET‐examination and metabolite evaluation in monkey of [11C]NAD‐299, a radioligand for visualisation of the 5‐HT1A receptor. Nucl Med Biol 29(1): 39-45. Sandell J, Halldin C, Hall H, et al. 1999. Radiosynthesis and autoradiographic evaluation of [C‐11]NAD‐299, a radioligand for visualization of the 5‐HT1A receptor. Nucl Med Biol 26: 159-164. Schou M, Halldin C, Pike VW, Mozley PD, Dobson D, et al. 2005. Post‐mortem human brain autoradiography of the norepinephrine transporter using (S,S)‐[18F]FMeNER‐D2. Eur Neuropsychopharmacol 15(5): 517-520. Schou M, Halldin C, Sovago J, Pike VW, Gulyas B, et al. 2003. Specific in vivo binding to the norepinephrine transporter demonstrated with the PET radioligand, (S,S)‐[11C] MeNER. Nucl Med Biol 30(7): 707-714. Schou M, Halldin C, Sovago J, Pike VW, Hall H, et al. 2004. PET evaluation of novel radiofluorinated reboxetine analogs as norepinephrine transporter probes in the monkey brain. Synapse 53(2): 57-67. Schou M, So´va´go´ J, Pike VW, Gulya´s B, Bogeso KP, et al. 2006. Synthesis and positron emission tomography evaluation of three norepinephrine transporter radioligands: [C‐11]desipramine, [C‐11]talopram and [C‐11]talsupram. Mol Imaging Biol 8(1): 1-8. Seneca N, Andree B, Sjoholm N, Schou M, Pauli S, et al. 2005. Whole‐body biodistribution, radiation dosimetry estimates for the PET norepinephrine transporter probe (S,S)‐[18F] FMeNER‐D2 in non‐human primates. Nucl Med Commun 26(8): 695-700. Sobrio F, Debruyne D, Dhilly M, Chazalviel L, Camsonne R, et al. 2005. Evaluation in rats and primates of [11C]‐mecamylamine, a potential nicotinic acetylcholine receptor radioligand for positron emission tomography. Neurochem Int 46(6): 479-488. Solin O, Eskola O, Hamill TG, Bergman J, Lehikoinen P, et al. 2004. Synthesis and characterization of a potent, selective, radiolabeled substance‐P antagonist for NK1 receptor quantitation: ([18F]SPA‐RQ). Mol Imaging Biol 6(6): 373-384. So´va´go´ J, Farde L, Halldin C, Langer O, Laszlovszky I, et al. 2004. Positron emission tomographic evaluation of the putative dopamine‐D3 receptor ligand, [11C]RGH‐1756 in the monkey brain. Neurochem Int 45(5): 609-617. So´va´go´ J, Farde L, Halldin C, Schukin E, Schou M, et al. 2005. Lack of effect of reserpine‐induced dopamine depletion on

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the binding of the dopamine‐D3 selective radioligand, [11C] RGH‐1756. Brain Res Bull 67(3): 219-224. Szabo´ Z, Kao PF, Mathews WB, Ravert HT, Musachio JL, et al. 1996. Positron emission tomography of 5‐HT reuptake sites in the human brain with C‐11 McN5652 extraction of characteristic images by artificial neural network analysis. Behav Brain Res 73(1–2): 221-224. Szabo´ Z, Ravert HT, Gozukara I, Geckle W, Seki C, et al. 1993. Noncompartmental and compartmental modeling of the kinetics of carbon‐11 labeled pyrilamine in the human brain. Synapse 15(4): 263-275. Szabo´ Z, Scheffel U, Suehiro M, Dannals RF, Kim SE, et al. 1995. Positron emission tomography of 5‐HT transporter sites in the baboon brain with [11C]McN5652. J Cereb Blood Flow Metab 15(5): 798-805. Takahashi K, Murakami M, Miura S, Iida H, Kanno I, et al. 1999. Synthesis and autoradiographic localization of muscarinic cholinergic antagonist (þ)N‐[11C]methyl‐3piperidyl benzylate as a potent radioligand for positron emission tomography. Appl Radiat Isot 5: 521-525. Tashiro M, Sakurada Y, Iwabuchi K, Mochizuki H, Kato M, et al. 2004. Central effects of fexofenadine and cetirizine: Measurement of psychomotor performance, subjective sleepiness, and brain histamine H1‐receptor occupancy using 11C‐doxepin positron emission tomography. J Clin Pharmacol 44(8): 890-900. Theodore WH, Carson RE, Andreasen P, et al. 1992. PET imaging of opiate receptor binding in human epilepsy using [F‐18]cyclofoxy. Epilepsy Res 13: 129-139. Tsukada H, Nishiyama S, Kakiuchi T, Ohba H, Sato K, et al. 2001a. Ketamine alters the availability of striatal dopamine transporter as measured by [11C]b‐CFT and [11C]b‐CIT‐FE in the monkey brain. Synapse 42(4): 273-280. Tsukada H, Takahashi K, Miura S, Nishiyama S, Kakiuchi T, 2001b. Evaluation of novel PET ligands (þ)N‐[11C]methyl‐ 3‐piperidyl benzilate ([11C](þ)3‐MPB) and its stereoisomer [11C](–)3‐MPB for muscarinic cholinergic receptors in the conscious monkey brain: A PET study in comparison with. Synapse 39(2): 182-192. Van Dort ME, Kim JH, Tluczek L, Wieland DM. 1997. Synthesis of 11C‐labeled desipramine and its metabolite 2‐hydroxydesipramine: Potential radiotracers for PET studies of the norepinephrine transporter. Nucl Med Biol 24(8): 707-711. Vas A, Gulya´s B. 2005. Eburnamine derivatives and the brain. Med Res Rev 25(6): 737-757. Volkow ND, Fowler JS, Ding YS, Wang GJ, Gatley SJ. 1998. Positron emission tomography radioligands for dopamine transporters and studies in human and nonhuman primates. Adv Pharmacol 42: 211-214. Volkow ND, Fowler JS, Ding YS, Wang GJ, Gatley SJ. 1999. Imaging the neurochemistry of nicotine actions: Studies

99

100

3

In vivo imaging of neurotransmitter systems with PET

with positron emission tomography. Nicotine Tob Res 1: S127-S132. Wagner HN, Burns HD, Dannals RF, et al. 1983. Imaging dopamine receptors in the human brain by positron tomography. Science 221: 1264-1266. Wienhard K, Coenen HH, Pawlik G, Rudolf J, Laufer P, et al. 1990. PET studies of dopamine receptor distribution using [18F]fluoroethylspiperone: Findings in disorders related to the dopaminergic system. J Neural Transm Gen Sect 81(3): 195-213. Wilson AA, Ginovart N, Hussey D, Meyer J, Houle S. 2002. In vitro and in vivo characterisation of [11C]‐DASB: A probe

for in vivo measurements of the serotonin transporter by positron emission tomography. Nucl Med Biol 29(5): 509-515. Yu M, Tueckmantel W, Wang X, Zhu A, Kozikowski AP, et al. 2005. Methoxyphenylethynyl, methoxypyridylethynyl and phenylethynyl derivatives of pyridine: Synthesis, radiolabeling and evaluation of new PET ligands for metabotropic glutamate subtype 5 receptors. Nucl Med Biol 32(6): 631-640. Zubieta JK, Koeppe RA, Frey KA, Kilbourn MR, Mangner T, et al. 2001. Assessment of muscarinic receptor concentrations in aging and Alzheimer disease with [11C]NMPB and PET. Synapse 39: 275-287.

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Synaptic and Nonsynaptic Release of Transmitters

E. S. Vizi . B. Lendvai

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Historical Background . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 102

2 Communication Between Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 103 2.1 Synaptic Interaction Between Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 103 2.2 Nonsynaptic Interaction Between Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 103 3 Release of Transmitters . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 103 3.1 Nonsynaptic Release of Transmitter . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 103 3.2 Spillover of Transmitters . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 105 4 4.1 4.2 4.3 4.4

What Influences the Concentration of Transmitter in the Extracellular Space? . . . . . . . . . . . . . . . 106 Amount of Transmitter Released . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 106 Uptake of Transmitters by Plasma Membrane Transporters . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 106 Volume of Extracellular Space . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 107 Effect of Drugs on Targets Located Intrasynaptically and Extrasynaptically: Law of Mass Action . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 107

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Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 108

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Abstract: Nonclassical receptorial functions represent revolutionary possibilities at the cellular level for some less-nderstood features of neural and cerebral activities. Although different forms of nonsynaptic communication often appear in different studies, their difference from synaptic actions is generally not recognized. The corner stones of interneuronal nonsynaptic communication include the release of transmitters into the extracellular space and the extrasynaptic receptors and transporters. Transmitters can be released from nonsynaptic varicosities without being coupled to frequency‐coded neuronal activity and from synapses following high presynaptic activity via spillover. The released substances are able to diffuse over large distances to reach remote tissue. Extrasynaptic receptors may occur at all possible membrane surfaces in various systems. These receptors are of high affinity, providing targets for low-dose drugs in many instances of medical therapy. List of Abbreviations: ACh, acetylcholine; AMPA, alpha-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid; CNS, central nervous system; DA, dopamine; 5-HT, serotonin; GABA, gamma-aminobutyric acid; NA, noradrenaline; NMDA, N-methyl-D-aspartate; nAChR, nicotinic acetylcholine receptor; VTA, ventral tegmental area

1

Historical Background

Of all the cells in the body, only nerve cells are able to communicate regularly with one another. In the nineteenth century, it was believed that ‘‘nerve centers’’ were made up of a continuous intermediary network between the motor nerves and the sensitive and sensory nerves (Cajal, 1937). Ramon y Cajal in his experiments applied Golgi staining to discover that neurons are independent units, and not entities fused to each other. This was an important step that changed the way of thinking of many scientists. Ramon y Cajal received the Nobel Prize in 1906 for this discovery. The question arose of how neurons communicate, if they do not fuse to one another with anastomosis, forming large neural nets of an intermediary network. Bernard demonstrated the dissociation of nerve and muscle activity by curarizing frogs (Bernard, 1857). This is regarded as one of the key experiments in the development of the concept of chemical transmission. The idea that nerves are able to release chemicals to communicate with other cells was first explicitly proposed for sympathetic nerves when Elliott, a young medical student in Cambridge, suggested in 1904, ‘‘adrenalin might. . .be the chemical stimulant liberated on each occasion when the impulse arrives at the periphery.’’ This brilliant hypothesis was confirmed by Loewi (1921), who showed that the stimulation of sympathetic nerves in frog heart is mediated by Acceleransstoff (adrenaline). The observation that the action of adrenaline and sympathetic stimulation are similar (Elliot, 1905) was further supported by the finding that the action of acetylcholine (ACh) and parasympathetic stimulation are also similar (Loewi, 1922; Dale, 1956). In spite of strong evidence suggesting otherwise, the alternative view that transmission is electrical enjoyed rather wide support during the first half of the twentieth century. This was mainly due to the fact that the electrical properties of what is now called conduction and transmission were seen to be similar (Erlanger, 1939; Gasser, 1939; Eccles, 1946). This belief gradually gained currency; a large number of neuroscientists believed that findings obtained in the neuromuscular junction were relevant not only to autonomic, but also to central synaptic transmission. Eccles in 1946 wrote ‘‘The original hypothesis was made as general as possible by applying it to the neuromuscular functions of skeletal muscle, and to the synapses of the sympathetic ganglia as well as of the central nervous system.’’ In fact, Eccles believed that the primary transmitter was electrical, but that chemical transmitters could be responsible only for slower and longer responses, detectable as a tail to the transmitter action on the postsynaptic site. This assumption was accepted almost universally by scientific society. Although different scientists (Dale, Feldberg, Kuffler, Uvna¨s, etc.) provided rather strong evidence to support and confirm the hypothesis of chemical transmission, Eccles resisted until 1948, when he acknowledged that even the fast response is due to acetylcholine at the neuromuscular junction and accepted the idea of chemical transmission. After his Pauline conversion from electrical to chemical transmission, the community of neuroscientists generally accepted the theory of chemical transmission for example, the communication between nerves and between nerve endings and target cells is chemical. It means that chemicals are released from nerve terminals in response to electrical depolarization followed by Ca2þ‐influx transmit messages between pre‐ and postsynaptic sites.

Synaptic and nonsynaptic release of transmitters

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Communication Between Cells

2.1 Synaptic Interaction Between Cells From Sherrington’s classical work on ‘‘Integrative action of the nervous system,’’ it has been generally accepted that the synapse, the ‘‘surface of separation’’ between neurons, is the primary site of neuronal information processing. Transmitter is released into the synaptic cleft in quantal packages. The average neuron forms about 1,000 synapses and it receives about 10,000 inputs. Since the human brain contains about 1011 nerve cells, it means that in the brain there are at least 1014 synapses for information processing. The generally accepted form of chemical communication between nerves and between nerve endings and target cells is that the transmitter is released into the synaptic cleft in quantal packages as a result of action potentials arriving at the terminals. The transmitter acts on receptors located on the postsynaptic site and either opens or closes ion channels, thereby establishing chemical communication between pre‐ and postsynaptic sites. The effect of transmitters is terminated by either enzymatic degradation (e.g., in case of ACh) or by active reuptake into nerve terminals by transporters (cf. Amara and Kuhar, 1993; Raiteri et al., 2002). Nevertheless, our current knowledge of how information is conveyed chemically from one cell to another is derived from and heavily influenced by the textbook data regarding the neuromuscular junction (cf. Katz, 1969), where the transmitter is released in quanta. This system is adopted for very fast signaling; the information transfer occurs within millisecond time intervals and is able to transmit messages at a rate of several hundred impulses per second.

2.2 Nonsynaptic Interaction Between Neurons In addition to transmitter substances acting at close range in chemical synaptic neurotransmission, chemical interaction exists and information processing occurs between neurons and between neurons and target cells without any close synaptic contact; there is a nonsynaptic communication system which operates over some distance in the extracellular spaces (cf. Vizi, 1974, 1979, 1980, 1984, 2000; Vizi et al., 1985; Agnati et al., 1986, 1995; Fuxe and Agnati, 1991; Bach‐y‐Rita, 1993; Vizi and Kiss, 1998). In the past few years, several neurochemical, anatomical, pharmacological, and neurophysiological observations have been made which suggest that chemical interaction between cells does not only take place across the synaptic gap between pre‐ and postsynaptic membranes but may also occur in the absence of such specialized contacts, i.e., nonsynaptically (> Figure 4-1). Neurochemical evidence has been obtained suggesting that noradrenaline (NA) released from axon terminals, which do not make synaptic contact with cholinergic terminals in the gut (Furness and Costa, 1974; Gordon‐Weeks, 1982), inhibits the release of acetylcholine from cholinergic varicosities of the Auerbach plexus (Vizi, 1968; Paton and Vizi, 1969; Knoll and Vizi, 1970; Vizi and Knoll, 1971). A very similar observation was first made in the cerebral cortex (Vizi, 1974, 1979, 1980) where the majority of noradrenergic varicosities do not make synaptic contacts (Descarries et al., 1977). This new concept of information processing, now known as nonsynaptic chemical transmission (Vizi, 1980, 1984), has been shown to be a rule rather than an exemption in the CNS (central nervous system) and has gained widespread acceptance (since 1986, it has also been called volume transmission, Agnati et al., 1986; Fuxe and Agnati, 1991; Agnati et al., 1995, paracrine release, spillover, nonconventional release, etc). Compelling neurochemical, functional, and pharmacological evidence (cf. Vizi, 2000) has accumulated suggesting that transmitters released from axon terminals are able to diffuse far away from the release site and have an effect on receptors located nonsynaptically.

3

Release of Transmitters

3.1 Nonsynaptic Release of Transmitter The idea that transmitters can be released from nonsynaptic areas was first suggested for transmitters in the autonomic nervous system, where the axon terminals rarely make synaptic contact with the target cells.

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. Figure 4-1 Nonsynaptic chemical transmission. A proportional diagram of two synaptic clefts: a typical synapse with a gap of 20 nm and a free axon terminal with its remote target cell (e.g., the vegetative nervous system). Note the difference in volume. Let us assume that both transmission sites are cholinergic. The average diameter of a vesicle containing acetylcholine (ACh) is 50 nm, therefore its volume is about 65,000 nm3 and its ACh concentration is about 0.1 M (cf. Marchbanks, 1979). If we assume that its content is completely discharged into a small synaptic cleft whose volume is about 200,000 nm3 (20 x 100 x 100 nm), the final concentration of ACh in the cleft is 30 mM (0.1/(200,000/65,000)), which is an extremely high concentration. Let us suppose that, for example, in Auerbach’s plexus, where the target smooth muscle cell is far from the varicose axon terminals (100–1,000 nm), only one vesicle is released. The volume in which the ACh released is 109 nm3 (1,000 x 1,000 x 1,000), i.e., a volume 104 times larger than that of a vesicle. Therefore, the ACh released from the vesicle is diluted by a factor of 10,000. If the cholinesterase is not active, the final concentration of ACh which reaches the muscarinic receptors of the smooth muscle is about 10
4 M. In this calculation, the cytoplasmic release has not been taken into account. Note the concentration of ACh in the cleft

Since then, a large body of evidence has shown that transmitters/modulators can be released from regions other than the nerve‐ending. Electron microscope and histochemical studies of the relationship between nerve terminals and target cells have shown that there are wide varieties of normal distances. The minimum width of the cleft between nerve varicosities and effector cells varies considerably in different tissues. In the vas deferens and sphincter pupillae, the separation is about 15–20nm. In blood vessels, the smallest space between varicosities in the perivascular plexus at the advential‐medial border and smooth muscle cells varies from about 50 nm to 2 mm (small muscular arteries, large arterioles, large elastic arteries) (see Burnstock, 1979). In these cases, no postjunctional specializations have been found with any consistency for wider neuromuscular junctions. Release from axonal varicosities devoid of synaptic membrane specialization has recently been suggested to be the function of the central monoamine terminals (Descarries et al., 1977; Beaudet and Descarries, 1978). Beaudet and Descarries (1978) claimed that the release of biogenic amines solely from varicosities making synaptic contact could hardly account for the total amount released from axon terminals. As the number of locus coeruleus cells in the rat is only about 1,400 (Descarries and Saucier, 1972) and the density of noradrenergic varicosities is 2 million/mm3, each cell body has an average of about 140,000 varicosities in the hippocampus. It means that the excitatory inputs to a noradrenergic cell body might activate a neuron whose transmitter, noradrenaline, released from varicosities might control a rather large field. It also means that a transmitter or modulator released from nonsynaptic varicosities could affect very large neuronal assemblies.

Synaptic and nonsynaptic release of transmitters

4

There is an interesting difference in the localization of receptors on effector cells where the transmitter is released into a small synaptic gap compared with those where it is released into a large extraneuronal space. In the former case, there is a small area of the cell where the receptors are concentrated. At the neuromuscular junction, for example the extrasynaptic area is relatively insensitive to the transmitter, because only a few receptors are there. However, when the transmitter release site and the target cells (e.g., Auerbach’s plexus varicose axon terminals and smooth muscle cells) are widely separated from each other (100–1,000 nm), there is no specific subsynaptic arrangement, and the receptors are evenly distributed along the whole surface of the smooth muscle cell. This morphological arrangement accommodates any type of diffusion‐mediated transmission, where the advantages of quantal release cannot be used. In the CNS, there are large amount of receptors located extrasynaptically and they are of high affinity. Even the cholinergic neurons fail to make synaptic contact in the hippocampus. Jones and Wonnacott (2004) provided evidence that in the ventral tegmental area (VTA), 27% of presynaptic a7 nAChRs (nicotinic acetylcholine receptors) are located extrasynaptically. The absence of axo‐axonic synapses (Descarries et al., 1997), i.e., direct cholinergic synaptic input to presynaptic a7 nAChRs, indicates that these receptors are likely to be activated by choline or ACh released from cholinergic varicosities (boutons) that are far away. Electron microscopy studies revealed that cholinergic varicosities in the hippocampal CA1 region are largely (93%) nonsynaptic compared with another transmitter system (Umbriaco et al., 1995), e.g., GABAergic and glutamatergic neurons, which make exclusively synaptic contacts. The nonsynaptic control of chemical neurotransmission by different modulators released from axonal varicosities lacking junctions might play a physiological role both in the CNS and in the neurovegetative system in shaping emotion, behavior, or learning processes, or in controlling the balance between the parasympathetic and the sympathetic nervous systems.

3.2 Spillover of Transmitters Though glutamate is the major excitatory transmitter, GABA is the most important inhibitory transmitter in the brain and spinal cord. Both glutamatergic and GABAergic terminals make exclusively synaptic contacts with other neurons. In this respect, they are different from monoaminergic nerve terminals which in the majority do not make synaptic contact (cf. Vizi, 2000). In recent years, it has become increasingly clear that receptors sensitive to glutamate and GABA besides their subsynaptic localization, they are also expressed extrasynaptically. The question that arises is where glutamate and GABA come from to signal these extrasynaptic receptors if they are only released into the synaptic cleft. The plausibility of the spillover (> Figure 4-2) depends on how much glutamate or GABA is released and how easily it can diffuse out of the synaptic cleft and how transporters terminate it to diffuse away. Glutamate, for example, the major excitatory transmitter of the brain, participates mainly in synaptic interactions between glutamatergic release sites predominantly located within synapses (Umbriaco et al., 1995) and AMPA and NMDA (N‐methyl‐D‐aspartate) receptors. Some synaptic spillover of glutamate has been observed and its effect on extrasynaptic NMDA (but not on AMPA) receptors was shown (Asztely et al., 1997; Kullmann and Asztely, 1998; Semyanov and Kullmann, 2000). The probability of spillover depends on how much glutamate is released, how easily it can diffuse out of the synaptic cleft, and how intra‐ and extrasynaptic transporters terminate it to diffuse away. The diffusion of glutamate away from synapses is therefore very limited because of effective neuronal and glial uptake processes. As far as the functional role of spillover is concerned, it has been shown (Mitchell and Silver, 2000) that spillover of glutamate released from excitatory mossy fibers is able to inhibit GABA release from neighboring Golgi cell terminals by activating presynaptic mGluRs. This heteroreceptor‐mediated inhibition of inhibitory fibers, in fact, boosts the efficacy of excitatory fibers. Kaneda and colleagues (1995) showed in cerebellar granule cells using voltage‐clamp experiments that in response to application of GABAA receptor antagonists there is a reduction in the ‘‘holding current.’’ A similar observation was made by Nusser and Mody (2002) on granule cells of the dentate gyrus. The inhibitory role of ambient GABA concentration was also shown (Semyanov et al., 2003, 2004) in hippocampal interneurons of CA1 region of the hippocampus, but interestingly not in adult pyramidal cells of hippocampus (Demarque et al., 2002). Nevertheless, there is a significant tonic GABAA

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. Figure 4-2 Spillover of synaptic transmitters in the central nervous system. In this scheme, a presynaptic axon terminal (dark gray) makes a synapse on a dendritic spine (light gray). Transmitters (black dots) in the vesicles are released into the synaptic cleft where it reaches the synaptic receptors (pentameric structures in this case). Arrows indicate the spillover of the released transmitters from the synapse

receptor mediated current in pyramidal cells when the GABA uptake is inhibited (Bai et al., 2001). During development, the tonic activation of GABAA receptors is induced by GABA released in a [Ca2þ]o‐dependent way, but in adult rats the tonic activation of GABAA receptors is produced by nonvesicular transmitter release (Rossi et al., 2003). This fact indicates that high‐affinity GABAA receptors are activated by extrasynaptic ambient GABA concentrations (> Figure 4-2).

4

What Influences the Concentration of Transmitter in the Extracellular Space?

4.1 Amount of Transmitter Released The amount of transmitter released is determined by the amplitude of the depolarization of the nerve terminals, which, in turn, is determined by the number and frequency of the action potentials in the axons. Transmitters are released in the form of packets, in quanta, that correspond to synaptic vesicles. There seems to be no evidence that quantal release can occur solely at varicosities with synaptic contact. In the varicosities of nerves, transmitter release occurs intermittently following the stimulation of the parent axon (Cunnane and Stja¨rne, 1982; Blakely et al., 1986) and is facilitated by high‐frequency train stimulation (Cunnane and Stja¨rne, 1984). There is a low probability of release in any varicosity invaded by a nerve action potential (Cunnane and Stja¨rne, 1982). The intermittency might be due to a failure of conduction of nerve action potentials within the varicose terminals (Stja¨rne, 1978; Morita and North, 1981) so that large parts of the distal region of arborization could be intermittently excluded from transmitter secretion. In places where the gap is large, where there is no synaptic specialization, or where the transmitter must cross distances of micrometers to reach the target cell, the transmitter released from the cytoplasm also plays a critical role in chemical transmission (Vizi et al., 1982). In fact, there is a large body of available evidence showing that the release of transmitter of cytoplasmic origin is also involved (carrier‐mediated release).

4.2 Uptake of Transmitters by Plasma Membrane Transporters Once a transmitter is released into extrasynaptic space, its effect on receptors is terminated by its reuptake into the surrounding nerve terminals and glia, a process mediated by plasma membrane transporters. These nonsynaptic transporters also terminate the overspill of the synaptically released transmitter, and thereby they play an important role in influencing the concentration of transmitters in the extraneuronal space. Monoamine uptake carriers belong to the family of Naþ/Cl
‐dependent membrane transporters containing

Synaptic and nonsynaptic release of transmitters

4

12 transmembrane domains (Amara and Arriza, 1993). The cloning and sequencing of monoamine transporters in the early 1990s (Blakely et al., 1991; Giros et al., 1991, 1992; Pacholczyk et al., 1991; Ramamoorthy et al., 1993) revealed that these proteins show a very high degree of structural homology. Extrasynaptic glutamate spillover was shown in the hippocampus (Asztely et al., 1997). It was also shown that glutamate transporters play a critical role in terminating the nonsynaptic diffusion of glutamate, thereby limiting cross talk between neighboring excitatory synapses. The activity of transporters is temperature‐ dependent (Amara and Arriza, 1993). Voltammetry and microdialysis techniques provided temporally resolved information concerning the concentration of transmitters in the extrasynaptic space. It turns out that in clinical practice, the chemicals are able to reach 0.1–23 mM in the extraneuronal space of the brain (> Table 4-1). . Table 4-1 A few examples of presynaptic autoreceptors and heteroreceptors able to inhibit or facilitate the release of neurotransmitters Neurotransmitter Acetylcholine Noradrenaline Dopamine 5‐HT GABA Glutamate

Inhibitory autoreceptor M2 a2A/D

Facilitatory autoreceptor nAChR b2

D2/D3 5‐HT1D GABAB Metabotropic, CB1

– 5‐HT – –

Inhibitory heteroreceptors a2, D2/D3, 5‐HT1B Opiate, H3, M2, D2, PGE2, GABAB M2 a2 GABAB, CB1, M2 –

Facilitatory heteroreceptors NMDA, nAChR Angiotensin II, nAChR, NMDA, GABAA, P2x7 nAChR, NMDA – – nAChR

Abbreviations: M2, muscarinic acetylcholine receptor; nAChR, nicotinic acetylcholine receptor; NMDA, N‐methyl‐D‐aspartate; PG, Prostaglandin. For literature, see Vizi and Kiss (1998), Starke (2001), Go¨bel et al. (2000)

The high degree of homology may explain the accumulating observations that functional segregation of monoaminergic pathways is not as marked as it was assumed previously. Accumulating evidence indicates the promiscuity of nonsynaptically located monoamine transporters. Using selective uptake blockers and specific pathway lesions, it was proved that (i) [3H]DA could be taken up by noradrenergic and serotonergic neurons (Descarries et al., 1987), (ii) dopaminergic terminals take up and release [3H]5‐HT in the striatum, (iii) serotonergic varicosities take up and release [3H]DA in the hippocampus of rabbit (Feuerstein et al., 1986), (iv) serotonergic transporters take up NA, and (v) serotonergic varicosities can release NA (Vizi et al., 2004). These findings may provide a better understanding of the functional properties of monoaminergic systems and the mechanism of action of antidepressant drugs.

4.3 Volume of Extracellular Space It has been shown that all the transmitters in the CNS are present in the extracellular space, which is about 12– 25% of the brain volume (Nicholson, 1985). This space has been called ‘‘communication channel’’ (Nicholson et al., 1979), because the migration of chemical signals by diffusion plays a very important role in nonsynaptic transmission. Tortuosity and volume fraction of the extracellular space modifies the diffusion (Nicholson, 2005).

4.4 Effect of Drugs on Targets Located Intrasynaptically and Extrasynaptically: Law of Mass Action According to pharmacological textbooks, most drugs produce their effects by binding to protein molecules (receptors, transporter molecules, enzymes, and ion channels). It is generally accepted that the magnitude of the biological response produced by an endogenous ligand is related to the number of receptors (target proteins)

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occupied (Rang, 2006); the receptor can bind only one drug molecule at a time. The first step in the action of drugs on target proteins (receptors) is the formation of a reversible drug‐target protein (receptor) complex. AðdrugÞ þ Rðfree receptorÞ ! ARðcomplexÞ: This step is governed by the Law of Mass Action; therefore, the actual concentrations of agonist (endogenous ligand, e.g., noradrenaline) and antagonist (e.g., drug applied) and the affinity of target proteins (receptors) play very important roles in the effect. Suppose that the intrasynaptic concentration of transmitter released into the synaptic cleft is between 1 and 10 mM, and that the orally administered drug (e.g., 10 mg/70 kg) can reach a concentration of about 0.05–5 mM in the extracellular space (> Table 4-2).

. Table 4-2 Concentration of drugs in the extracellular space Concentration (mM) Drug

Plasma

Nicotine Imipramine Citalopram Desmethylimipramine Fluoxetine

0.4–4.5 1–2 1 2 1

Cerebrospinal fluid

0.1

References Zevin et al. (1998) Besret et al. (1996) Hyttel (1982) Muscettola et al. (1978) Pato et al. (1991)

It is expected that a similar concentration would be found in the synapse. Therefore, taking into account the Law of Mass Action, the drug effect is marginal, if any. Therefore, the site of action of most drugs is those binding proteins (receptors, transporters, ion channels, and enzymes), which are located nonsynaptically, i.e., outside the synapse, in the extracellular space where the receptors and transporters are of high affinity (Vizi, 2000).

5

Conclusions

The nonsynaptic control of chemical neurotransmission by different modulators released from axonal varicosities lacking junctions might play a physiological role both in the CNS and in the neurovegetative system in shaping emotion, behavior, or learning processes, or in controlling the balance between the parasympathetic and sympathetic nervous system. Are the receptors located outside the postsynaptic density of the synapse functionally a part of chemical transmission, or are they promiscuous and accessible to chemicals released from different boutons with (in case of spillover) and without synaptic arrangements? The answer is yes. Presynaptic release modulating receptors represent suitable targets for pharmacological intervention by exogenous compounds acting as agonists, partial agonists, or antagonists. It is thus possible that presynaptic release modulating autoreceptors and heteroreceptors (> Figure 4-3) of high affinity may become the target of action for a new generation of drugs which can produce the desired therapeutic actions through the modulation or fine tuning of the release of neurotransmitters or cotransmitters. This novel mechanism differs from the well‐ established approach of using agonists or antagonists to directly stimulate or block postsynaptic receptors. An important way to control the activation of nonsynaptic receptors is to regulate the levels of transmitters in the extracellular space. Plasma membrane transporters play a very important role in terminating the levels of the transmitters released into the extraneuronal space. Clinically applied antidepressants reaching concentrations of 0.5–13 mM in the extraneuronal space may exert their effects on high‐affinity nonsynaptic transporters (Vizi, 2000). Because of the major impact of tonic inhibition or stimulation by endogenous transmitters on neuronal activity, this form of influence, besides its physiological importance, could be an

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. Figure 4-3 Role of autoreceptors and heteroreceptors in modulation of transmitter release evoked by neuronal activity. This type of release is [Ca2+]‐dependent. Heteroreceptor is sensitive to a transmitter which is not produced by the neuron on which the receptor is expressed. Autoreceptors are the receptors sensitive to the neurons’ own transmitter substance. Scheme shows an example: noradrenaline (NA) inhibits its own release via the stimulation of presynaptic a2‐autoreceptors. Noradrenaline inhibits the release of acetylcholine (ACh) via the activation of presynaptic a2‐heteroreceptors. The effect of NA is terminated by its reuptake

important novel pharmacological target for the treatment of a wide range of disorders. Nonsynaptic and high‐ affinity GABAA receptors responsible for the tonic inhibitory conductance may be of clinical importance as targets for anesthetics and sedative drugs (Bai et al., 2001). It seems very likely that GABA and glutamate receptors need not be restricted to synapses to serve physiological functions.

References Agnati LF, Zoli M, Stromberg I, Fuxe K. 1995. Intercellular communication in the brain: Wiring versus volume transmission. Neuroscience 69: 711-726. Agnati LF, Fuxe K, Zoli M, Ozini I, Toffano G, et al. 1986. A correlation analysis of the regional distribution of central enkephalin and beta‐endorphin immunoreactive terminals and of opiate receptors in adult and old male rats. Evidence for the existence of two main types of communication in the central nervous system: The volume transmission and the wiring transmission. Acta Physiol Scand 128: 201-207. Amara SG, Arriza JL. 1993. Neurotransmitter transporters three distinct gene families. Curr Opin Neurobiol 3: 337-344. Amara SG, Kuhar MJ. 1993. Neurotransmitters: Recent progress. Annu Res Neurosci 16: 73-93. Asztely F, Erdemli G, Kullmann DM. 1997. Extrasynaptic glutamate spillover in the hippocampus: Dependence on temperature and the role of active glutamate uptake. Neuron 18: 281-293. Bach‐y‐Rita P. 1993. Neurotransmission in the brain by diffusion through the extracellular fluid: A review. Neuroreport 4: 343-350. Bai D, Zhu G, Pennefather P, Jackson MF, MacDonald JF, Orser BA. 2001. Distinct functional and pharmacological properties of tonic and quantal inhibitory postsynaptic currents mediated by gamma‐aminobutyric acid (A)

receptors in hippocampal neurons. Mol Pharmacol 59: 814-824. Beaudet A, Descarries L. 1978. The monoamine innervation of rat cerebral cortex: Synaptic and non‐synaptic axon terminals. Neuroscience 3: 851-860. Bernard C. 1857. Lecons sur les Effects des Substances Toxiques et Medicamenteuses. Paris: Bailliere; pp. 351-353. Blakeley AGH, Cunnane TC, Muir TC. 1986. Effects of calcium channel antagonists on action potential conduction and transmitter release in the guinea‐pig vas deferens. Br J Pharmacol 89: 235-244. Blakely RD, Berson HE, Fremeau RT, Jr Caron MG, Peek MM, et al. 1991. Cloning and expression of a functional serotonin transporter from rat brain. Nature 354: 66-70. Burnstock G. 1979. The ultrastructure of autonomic cholinergic nerves and Junctions. Prog Brain Res 79: 3-21. Cajal SR. 1937. Recollections of my life. Philadelphia: American Philosophical Society. Cunnane TC, Stja¨rne L. 1982. Secretion of transmitter from individual varicosities of guinea‐pig and mouse vas deferens. J Physiol 399: 607-632. Cunnane TC, Stja¨rne L. 1984. Frequency dependent intermittency and ionic basis of impulse conduction in postganglionic sympathetic fibres of guinea‐pig deferens. Neuroscience 11: 211-229.

109

110

4

Synaptic and nonsynaptic release of transmitters

Dale HH. 1956. Published in, Nobel Lectures in Physiology or Medicine, 1922–1941. Elsevier; Amsterdam: pp. 402-413. Demarque M, Represa A, Becq H, Khalilov I, Ben‐Ari Y, Aniksztejn L. 2002. Paracrine intercellular communicationby a Ca2þ and SNARE‐independent release of GABA and Glutamate prior to synapse formation. Neuron 36: 1051-1061. Descarries L, Saucier G. 1972. Disappearance of the locus coeruleus in the rat after intraventricular 6‐hydroxdopamine. Brain Res 37: 310-316. Descarries L, Gisiger V, Steriade M. 1997. Diffuse transmission by acetylcholine in the CNS. Prog Neurobiol 70(3): 603-625. Descarries L, Watkins KC, Lapierre Y. 1977. Noradrenergic axon terminals in the cerebral cortex of the rat. III Topometric ultrastructural analyses. Brain Res 133: 197-222. Descarries L, Lemay B, Doucet G, Berger B. 1987. Regional and laminar density of the dopamine innervation in adult rat cerebral cortex. Brain Res 441: 807-824. Eccles JC. 1946. An electrical hypothesis of synaptic and neuromuscular transmission. Ann NY Acad Sci 47: 429-455. Elliot TR. 1905. The action of adrenaline. J Physiol Lond 32: 401-467. Erlanger J. 1939. The initiation of impulses in axons. J Neurophysiol 2: 370-379. Feuerstein TJ, Hertting G, Lupp A, Neufang B. 1986. False labeling of dopaminergic terminals in the rabbit caudate nucleus: Uptake and release of [3H]‐5hydroxytryptamine. Br J Pharmacol 88: 677-684. Furness JB, Costa M. 1974. The adrenergic innervation of the gastrointestinal tract. Rev Physiol 69: 1-51. Fuxe K, Agnati LF. 1991. Two principal modes of electrochemical communication in the brain: Volume versus wiring transmission. Volume transmission in the brain: Novel mechanisms for neural transmission. Fuxe K, Agnati LF, editors. New York: Raven Press; pp. 1‐10. Gasser HS. 1939. Axons as samples of nervous tissue. J Neurophysiol 2: 361-369. Giros B, el Mestikawy S, Bertrand L, Caron MG. 1991. Cloning and functional characterization of a cocaine‐sensitive dopamine transporter. FEBS Lett 295: 149-154. Giros B, el Mestikawy S, Godinot N, Zheng K, Han H, et al. 1992. Cloning, pharmacological characterization, and chromosome assignment of the human dopamine transporter. Mol Pharmacol 42: 383-390. Gordon‐Weeks PR. 1982. Noradrenergic and non‐noradrenergic nerves containing small granular vesicles in Auerbach’s plexus of the guinea pig. Neuroscience 11: 2925-2936. Jones IW, Wonnacott S. 2004. Precise localization of alpha 7 nicotinic acetylcholine receptors on glutamatergic axon terminals in the rat ventral tegmental area. J Neurosci 24: 11244-11252.

Kaneda M, Farrant M, Cull‐Candy SG. 1995. Whole‐cell and single channel currents activated by GABA and glycine in granule cells of the rat cerebellum. J Physiol 485: 419-435. Katz B. 1969. The release of neural transmitter substances, Sherrington lecture. Liverpool: Liverpool University Press. Knoll J, Vizi ES. 1970. Presynaptic inhibition of acetylcholine release by endogeneous and exogeneous noradrenaline at high rate of stimulation. Br J Pharmacol 40: 554-555. Kullmann DM, Asztely F. 1998. Extrasynaptic glutamate spillover in the hippocampus: Evidence and implications. Trends Neurosci 21: 8-14. ¨ ber humorale U ¨ bertragbarkeit der Loewi O. 1921. U Herznervenwirkung. Pflu¨gers Arch 189: 239-242. ¨ ber humorale U ¨ bertragbarkeit der Loewi O. 1922. U Herznervenwirkung II. Mitteilung, Pflugers Arch 193: 201-213. Mitchell SJ, Silver RA. 2000. Glutamate spillover suppresses inhibition by activating presynaptic mGluRs. Nature 404: 498-502. Morita K, North RA. 1981. Opiates and encephalin reduce the excitability of neuronal processes. Neuroscience 6: 1943-1951. Nicholson C. 1985. Diffusion from an injected volume of a substance in brain tissue with arbitrary volume fraction and tortuosity. Brain Res 333: 325-329. Nicholson C. 2005. Factors governing diffusing molecular signals in brain extracellular space. J Neural Transm 112: 29-44. Nicholson C, Phillips JM, Gardner‐Medwin AR. 1979. Diffusion from an iontophoretic point source in the brain: Role of tortuosity and volume fraction. Brain Res 169: 580-584. Nusser Z, Mody I. 2002. Selective modulation of tonic and phasic inhibitions in dentate gyrus granule cells. J Neurophysiol 87: 2624-2628. Pacholczyk T, Blakely RD, Amara SG. 1991. Expression cloning of a cocaine‐ and antidepressant‐sensitive human noradrenaline transporter. Nature 350: 350-354. Paton WDM, Vizi ES. 1969. Non‐synaptic modulation of transmitter release: Pharmacological implication. Br J Pharmacol 35: 10-28. Raiteri L, Raiteri M, Bonanno G. 2002. Coexistence and function of different neurotransmitter transporters in the plasma membrane of CNS neurons. Prog Neurobiol 68: 287-309. Ramamoorthy S, Bauman AL, Moore KR, Han H, Yang‐Feng T, et al. 1993. Antidepressant‐ and cocaine‐sensitive human serotonin transporter. Molecular cloning, expression, and chromosomal localization. Proc Natl Acad Sci USA 90: 2542-2546. Rang HP. 2006. The receptor concept: Pharmacology’s big idea. Br J Pharmacol 147 (Suppl 1): S9-S16.

Synaptic and nonsynaptic release of transmitters Rossi DJ, Hamann M, Attwell D. 2003. Multiple modes of GABAergic inhibition of rat cerebellar granule cells. J Physiol 548: 97-110. Semyanov A, Kullmann DM. 2000. Modulation of GABAergic signaling among interneurons by metabotropic glutamate receptors. Neuron 25: 663-672. Semyanov A, Walker MC, Kullmann DM. 2003. GABA uptake regulates cortical excitability via cell type‐specific tonic inhibition. Nat Neurosci 6: 484-490. Semyanov A, Walker MC, Kullmann DM, Silver RA. 2004. Tonically active GABA A receptors: Modulating gain and maintaining the tone. Trends Neurosci 34(2): 262-269. Stja¨rne L. 1978. Facilitation and receptor mediated regulation of noradrenaline secretion by control of recruitment of varicosities as well as by control of electrosecretory coupling. Neuroscience 3: 1147-1155. Umbriaco D, Garcia S, Beaulieu C, Descarries L. 1995. Relational features of acetylcholine, noradrenaline, serotonin and GABA axon terminals in the stratum radiatum of adult rat hippocampus (CA1). Hippocampus 5: 605-620. Vizi ES. 1968. The inhibitory action of noradrenaline and adrenaline on release of acetylcholine from guinea‐pig ileum longitudinal strips. Arch Exp Path Pharmacol 259: 199-200. Vizi ES. 1974. Interaction between adrenergic and cholinergic systems presynaptic inhibitory effect of noradrenaline on: Acetylcholine release. J Neural Transm Suppl 11: 61-78. Vizi ES. 1979. Presynaptic modulation of neurochemical transmission. Prog Neurobiol 12: 181-290.

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Vizi ES. 1980. Non‐synaptic modulation of transmitter release: Pharmacological implication. Trends Pharmacol Sci 1: 172-175. Vizi ES. 1984. Non‐synaptic interactions between neurons: Modulation of neurochemical transmission. London: Wiley. Vizi ES. 2000. Role of high‐affinity receptors and membrane transporters in nonsynaptic communication and drug action in the central nervous system. Pharmacol Rev 52: 63-90. Vizi ES, Kiss JP. 1998. Neurochemistry and pharmacology of the major hippocampal transmitter systems: Synaptic and nonsynaptic interactions. Hippocampus 8: 566-607. Vizi ES, Knoll J. 1971. The effects of sympathetic nerve stimulation and guanethidine on parasympathetic neuroeffector transmission: The inhibition of acetylcholine release. J Pharm Pharmacol 23: 918-925. Vizi ES, Ha´rsing LG Jr, Zima´nyi I, Gaa´l G. 1985. Release and turnover of noradrenaline in isolated median eminence: Lack of negative feedback modulation. Neuroscience 16: 907-916. Vizi ES, Zsilla G, Caron MG, Kiss JP. 2004. Uptake and release of norepinephrine by serotonergic terminals in norepinephrine transporter knock‐out mice: Implications for the action of selective serotonin reuptake inhibitors. J Neurosci 24: 7888-7994. Vizi ES, To¨ro¨k T, Seregi A, Serfo˝zo˝ p, Adam‐Vizi V. 1982. Na– K‐activated ATPase and the release of acetylcholine and noradrenaline. J Physiol (Paris) 78: 399-406.

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B. Lendvai

1

Historical Background . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 114

2

Synthesis, Storage, and Release of Acetylcholine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 114

3

Breakdown of ACh . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 115

4

Structure of the Central Cholinergic System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 116

5

Synaptic Versus Nonsynaptic Release of ACh . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 116

6 6.1 6.2 6.3 6.4 6.5 6.6 6.7 6.8 6.9

Nicotinic ACh Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 117 Subunits and Subtypes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 117 Selective Nicotinic Ligands . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 117 Movement of Ions After nAChR Activation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 117 Desensitization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 117 Presynaptic Nicotinic Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 118 Postsynaptic Nicotinic Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 118 Role of Nicotinic Receptors in Reward Mechanisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 119 Special Role of Nicotinic Receptors in Neural Plasticity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 119 Nicotinic Receptors in Synaptic and Nonsynaptic Transmission . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 120

7 7.1 7.2 7.3 7.4

Muscarinic ACh Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 120 Subunits and Subtypes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 120 Subcellular Action Mechanisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 121 Pharmacology of Muscarinic Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 122 Muscarinic Receptor Functions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 122

8

Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 124

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2008 Springer ScienceþBusiness Media, LLC.

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Abstract: The cholinergic system can modulate cognitive functions efficiently in the brain acting on a rich assembly of metabotropic and ionotropic receptors. The cholinergic system operates through the cooperation of the muscarinic and the nicotinic subsystems. While muscarinic ACh receptors mediate slow responses with considerable delay, nicotinic facilitation, following activation of nicotinic ACh receptors, evokes relatively fast responses. In some cases muscarinic and nicotinic ACh receptors form a dual control striatal on certain cell types, such as the spiny interneurons of the striatum. On important aspect of nicotinic transmission is that it modulates, rather than mediate, fast synaptic transmission. Desensitization of these receptors leads to a loss of function that is a key factor in the effect of nicotine during smoking. Desensitization extends the possible states of cholinergic transmission and increases the computational power of the neuron. As most nicotinic receptors are found in nonsynaptic localizations, especially on axons. They can directly release transmitters from presynaptic boutons. Importance of studies on nicotinic and muscarinic effects is highlighted by the fact that cholinergic therapy is the mainstay treatment for Alzheimer’s disease. The current view that nonsynaptic communication is dominant in cholinergic transmission also support the future perspective of drug therapy targeting high affinity nonsynaptic receptors. List of Abbreviations: ACh, acetylcholine; AChE, acetylcholine-esterase; ChAT, choline-acetyltranspherase; CNS, central nervous system; GABA, Gamma-aminobutyric acid; mAChR, muscarinic acetylcholine receptors; nAChR, nicotinic acetylcholine receptors; NMDA, N-methyl-D-aspartate

1

Historical Background

Cholinergic transmission has a unique position in the mind of neurochemists as being the first identified neurotransmitter during the first half of the twentieth century based on studies of Henry Dale, Otto Loewi, Feldberg and others. The very first discovery was the observation that a chemical substance (termed vagusstoff) is linked to the vagus action in the autonomic nervous system. Nicotinic receptors were the first receptors to be named. Today, studies focusing on the cholinergic system of the brain have received particular attention as the loss of cholinergic function is thought to underlie the age‐related learning impairments and memory loss that accompanies Alzheimer’s disease, one of the major health care problems in the world.

2

Synthesis, Storage, and Release of Acetylcholine

Acetylcholine (ACh) is synthesized in the boutons of cholinergic axons. Choline is taken up from the extracellular space by its specific transporter (> Figure 5-1). Although low‐affinity choline uptake is present in most tissues, cholinergic cells are equipped with a high‐affinity (Km
1–5 mM) choline uptake that provides stable choline supply under physiological conditions (choline is present in the plasma at about 10 mM). Hemicholinium, a plasma membrane choline transport inhibitor, causes disruption of ACh release especially during prolonged stimulation. Most of the choline derives from recycling of released ACh following hydrolysis. Another important source of choline is the breakdown of phosphatidylcholine. ACh is synthesized by the choline‐acetyltranspherase (ChAT), which transfers an acetyl group from acetyl coenzyme A to choline. Brain ChAT has a KD for choline of approximately 1 mM and a KD for acetyl coenzyme A of approximately 10 mM. There is an excess capacity of ChAT to produce ACh; the in vitro activity of the enzyme was found higher than that in vivo. The rate‐limiting step for ACh synthesis is likely the transport of acetyl coenzyme A, which comes from the inner membrane of mitochondria following the glucose– pyruvate transformation. The newly synthesized ACh is transported from the soma to the vesicles in the cholinergic axon terminal where it is stored. The transport of ACh can be prevented by incubation with vesamicol, which induces ACh accumulation in the plasma and the depletion of the vesicle pool of ACh. Most ACh molecules in the synaptosome, a neuronal preparation containing only the axon terminals, have been found associated with vesicles, as revealed by electron microscopy. ACh is typically released to the extracellular space from these vesicles following a fusion with the plasma membrane of the cell in

Cholinergic transmission

5

. Figure 5-1 Synthesis and nonsynaptic release of ACh. Cholinergic innervation arises from different loci where the cell body of the cholinergic projection neuron is located. Most of the ACh is released to the nonsynaptic, extracellular space (arrows). In contrast, ACh can be released in a few synaptic specializations for cholinergic transmission where ACh activates postsynaptic cell (gray shaded) in a one‐to‐one manner (small circle). Excitation of the postsynaptic cell, either by nonsynaptically or synaptically released ACh, occurs after activation postsynaptic/ dendritic nicotinic or muscarinic receptors. ACh is synthesized in the axon terminals where the high‐affinity choline uptake and the acetyl coenzyme A production provide the source molecules by ChAT. AChE is responsible for the breakdown of the released ACh

nonsynaptic sites (> Figure 5-1). From the functional aspect, it seems there are two distinct pools of ACh within the terminal: one ‘‘readily available’’ pool and a ‘‘reserve’’ or ‘‘depot’’ pool of ACh.

3

Breakdown of ACh

In the extracellular space, acetylcholine‐esterase (AChE) is responsible for the elimination of the effect of ACh by degrading the released ACh. AChE exists in several molecular forms (monomers, dimers, tetramers) with different solubility and subcellular localization. Cholinesterase genes encode a peptide without obvious membrane‐spanning regions making them available for secretion. Indeed, the soluble form of AChE can be released from nerve terminals. AChE shares structural similarity with neuroligins. During development the level of AChE regulates synaptogenesis for glutamatergic transmission (Dong et al., 2004). The distribution of AChE is relatively uniform in the central nervous system (CNS); it can be found in the cytoplasm and in the extracellular space even at large distances from cholinergic boutons. There are various ways to influence cholinergic transmission by modulating ACh release. Inhibitors of AChE, called anticholinesterases, induce accumulation of ACh in the extracellular space, and therefore prolong the action of the released ACh at the receptor. As a result, the decay of the postsynaptic current or potential by the released ACh is prolonged from 1–2 to 5–30 msec. On the system level, fasciculation and muscle twitching are initially observed after AChE inhibition, followed by flaccid paralysis. Some reversible inhibitors, such as gallamine and propidium, bind to a peripherial site, while other reversible inhibitors (neostigmine and physostigmine) act at the active site with an approximately 4‐h duration. The latter two inhibitors are used to treat glaucoma, myasthenia gravis, and dysfunction of smooth muscle. Another important application area of anticholinesterases is the pharmacotherapy of Alzheimer’s disease. Tacrine and donezepil are used to enhance cholinergic transmission in Alzheimer’s patients. Irreversible inhibitors completely inhibit the

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breakdown of ACh, and therefore are dangerous for life. These molecules are used as insecticides. Nerve gases (sarin, soman) can cause death within 5 min of exposure. On the receptive side of cholinergic transmission there are two major receptor types: nicotinic (nAChRs) and muscarinic ACh receptors (mAChRs). All subtypes of nAChRs mediate excitatory conductances through the receptor ion channel. Muscarinic receptors have metabotropic functions and they can be excitatory or inhibitory depending on the subtype and the downstream subcellular mechanism.

4

Structure of the Central Cholinergic System

The cholinergic system is built up by two main cell types: cholinergic interneurons and projection neurons. ACh‐containing interneurons occur in the striatum, where the large aspiny neurons provide the cholinergic tone of the striatum. These cells are under the tonic inhibition of the nigrostriatal dopaminergic system through D2 receptors. In Parkinson’s disease the loss of this inhibitory influence results in high ACh concentration in the striatum. Prevention of the effect of ACh in Parkinson’s disease is an important part of the clinical therapy. Cholinergic interneurons were also identified in the hippocampus. Projection neurons include the cholinergic neurons of the medial septum, which provide an important innervation to the hippocampus, and the cholinergic neurons of the Meynert nucleus in the basal forebrain. The cholinergic neurons of the Meynert nucleus innervate the cerebral cortex. The cerebral cortex is diffusely innervated by cholinergic axons from the basal forebrain and axons from cholinergic interneurons (Eckenstein and Baughman, 1984). Cholinergic neurons of the medial habenula project to the interpeduncular nucleus. There are some other less investigated cholinergic loci in the CNS: vertical nucleus of the diagonal band projects to the hippocampus, neurons in the horizontal limb of the diagonal band innervate the olfactory bulb, the midbrain pedunculopontine and laterodorsal tegmental nucleus gives cholinergic input to the thalamus, and the cholinergic cells in the parabigeminal nucleus, which provide input to the superior colliculus.

5

Synaptic Versus Nonsynaptic Release of ACh

Ultrastructural morphometric studies demonstrated that the normal cholinergic innervation of adult rat parietal cortex (Umbriaco et al., 1994) and hippocampus (Umbriaco et al., 1995) predominantly does not result in synapses on other neurons (>85% of varicosities are without synaptic contact). These observations supported the idea that ACh participates primarily in nonsynaptic interactions (Descarries and Mechawar, 2000; Vizi, 2000). Cholinergic axons can be identified by staining for vesicular ACh transporter immunoreactivity, providing the strongest labeling in the stratum oriens of the CA1 region within the hippocampus (Towart et al., 2003). This observation indicates that the cholinergic input predominantly targets the basal dendrites of pyramidal neurons and the local interneurons. Another important evidence for the nonsynaptic nature of the cholinergic system is the somewhat surprising localization of AChE. The fact that AChE can be found in distant areas from cholinergic axon terminals and not restricted to the area of cholinergic release sites strongly suggests that the enzyme must sense and degrade ACh molecules that have traveled far in the extracellular space. This assumption is consistent with the nonsynaptic nature of most cholinergic boutons. ACh, released from the terminals without synaptic content, diffuses in the extracellular space to reach remote receptors where extracellular AChE terminates the cholinergic action. Neostigmine, a cholinesterase inhibitor, also enhances the extracellular level of noradrenaline measured by microdialysis (Kiss et al., 1999). During microdialysis, the samples are taken from the extracellular space, therefore, this method provides data for the nonsynaptic release of transmitters. It seems ACh is released from cholinergic boutons tonically; neostriatal cholinergic interneurons produce spontaneous tonic firing in the absence of synaptic input that results in a tonic release of ACh (Bennett and Wilson, 1999). The released ACh keeps the striatal DA terminals under a tonic control (Zhou et al., 2001). The release process can be pharmacologically manipulated; botulinum toxin A and tetanus toxin are known to block the release of ACh causing paralysis.

Cholinergic transmission

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Nicotinic ACh Receptors

6.1 Subunits and Subtypes Genes encoding neural nAChR subunits express nine a (a2–a10) and three b subunits (b2–b4), which can coassemble to form pentameric functional receptors (Role and Berg, 1996). Pharmacological studies on transgenic mice have shown that the different subunits vary in their distribution and channel properties. The a4b2 and the a7 subtypes are predominant in the CNS while the a3b4 subunit‐containing nAChR is predominant in the peripherial nervous system (Role and Berg, 1996). Because of the diversity of subunits and minimal requirement of five subunits to form the channel of nAChRs, one cannot predict nAChR compositions based solely on the set of genes expressed by the neuron. There is evidence indicating the existence of multiple functional subtypes of nAChRs in the same neuron. For example, activation of nAChRs on hippocampal stratum radiatum interneurons involves both a7 and non‐a7 subunits and causes depolarization and action potential generation (Frazier et al., 1998b; McQuiston and Madison, 1999).

6.2 Selective Nicotinic Ligands There are a few selective agonists for nAChRs. Choline seems to be a selective activator of a7 nAChRs and a partial agonist on the a3b4 nAChRs (Alkondon et al., 1997). The antagonists methyllycaconitine (MLA) and a‐bungarotoxin selectively block the homomeric a7 nAChRs. At higher concentration (mM), MLA blocks other nAChRs, such as a4b2. Mecamylamine is a non‐selective channel blocker of all nAChRs with the highest potency on a4b4 and with relatively weak potency at a7 receptors. Nicotine‐induced increase in mEPSC frequency can be fully antagonized by a‐bungarotoxin, while mecamylamine causes only a 70% inhibition (Sharma and Vijayaraghavan, 2003). Mecamylamine, at higher (100 mM) concentration, can block other ligand‐gated ion channels, such as N‐methyl‐D‐aspartate (NMDA) receptors. The antagonist dihydro‐b‐ erithroidine (DHbE) has relatively low affinity for the a7 nAChRs and very high affinity to the a4bb2 and the a4b4 nAChRs.

6.3 Movement of Ions After nAChR Activation Nicotinic receptors are highly permeable to Naþ, Kþ, and Ca2þ. The Ca2þ/Naþ permeability ratio for the a7 receptor is >10 (the average ratio for all nAChRs is >1) (Seguela et al., 1993). In general, central nAChRs differ from the muscle nAChRs receptors in that the neuronal types are more permeable to Ca2þ (Vernino et al., 1992). Indeed, in hippocampal interneurons and cerebellar granule cells, nAChR stimulation induces Ca2þ transients via a7 nAChRs (Didier et al., 1995; Khiroug et al., 2003). Although nicotine can indeed cause large influx of Ca2þ into the cell, it is usually corroborated by various amplification mechanisms including voltage‐sensitive Ca2þ channels (VSCCs) and intracellular Ca2þ stores. In rat chromaffin cells, a part of the nAChR stimulation‐evoked Ca2þ response is mediated by VSCCs (Khiroug et al., 1997). Voltage‐sensitive Naþ channels can amplify the action of VSCCs in different tissue preparations (Mulle et al., 1992; Vijayaraghavan et al., 1992; Soliakov and Wonnacott, 1996). Nevertheless, in certain cells, Ca2þ rises by nicotinic stimulation can be exclusively mediated by the nAChR ion channel.

6.4 Desensitization Resting nAChR channels open in response to agonist binding to allow passage of ions. Prolonged presence of an agonist produces a desensitized state that no longer permits ion movements through the channel. Recovery from this state occurs after the agonist has been removed. For a long time, the main reason for the failures to detect nAChRs has been the rapid desensitization. Ca2þ accumulation due to receptor activation takes longer and so it was somewhat easier to detect rises in intracellular free Ca2þ; that is why most of the

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first reports on cellular functions of central nAChRs emerged from optical studies (Vijayaraghavan et al., 1992). In outside‐out membrane patches of acutely isolated habenula neurons, applications of 100 mM nicotine produces macroscopic currents due to the opening of a large number of channels. During the continuous perfusion of the agonist, the number of open nAChR channels decreases exponentially because of receptor desensitization. A progressive loss in the number of channels with time is considered as receptor rundown (Lester and Dani, 1994). Desensitization expands the variability of nAChR‐mediated functions providing an extra computational power of the neuron. The findings that nicotine can cause similar effects to perfusion with nicotinic antagonists suggest a physiological role of desensitization of nAChRs during cigarette smoking (Chiodini et al., 1999; Zhou et al., 2001). Desensitization varies across cell types. In neurons, desensitization of nicotine‐induced currents becomes complete within a few seconds. Another important feature of the nicotinic desensitization is that the size of the intracellular Ca2þ transients can determine the rate of nAChR desensitization. In addition to the rapid desensitization, there is a long‐lasting or persistent inactivation of nAChRs, which, in contrast to desensitization, occurs immediately after the drug application and remains from 60 minutes to several hours (Lukas, 1991; Rowell and Duggan, 1998). The rates of recovery from desensitization depend on the time of agonist exposure and on the amount of agonist used to induce desensitization. The recovery time is about 10–30 sec after desensitization of nAChRs.

6.5 Presynaptic Nicotinic Receptors Presynaptic nAChRs have been described in various brain regions. These receptors that reside on the axon varicosity but far from the release site have been assigned the term preterminal receptors (Wonnacott, 1997). Why presynaptic (preterminal) nAChRs are so important for understanding cholinergic transmission? Activation of presynaptic nAChRs that precedes or coincides the arrival of an action potential into the terminal region of the neuron can increase the probability of release via the integrative capabilities of nAChR‐induced Ca2þ influx. The releasing action of presynaptic nAChRs are supported by several lines of experimental evidence: (1) the enhancement of the frequency and the size of excitatory synaptic potentials (EPSP) by nicotinic agonists leads to synchronization of the release process; this increase in the activity is sufficient to drive the postsynaptic cell above the firing threshold (Sharma and Vijayaraghavan, 2003) and (2) biochemical measurement of extracellular level of transmitters revealed that presynaptic nAChRs play an important role in regulating the release of different neurotransmitters (Vizi et al., 1995; Lendvai et al., 1996; Sershen et al., 1997; Vizi and Lendvai, 1999; Kofalvi et al., 2000). Different subtypes of nAChRs can be involved in the presynaptic regulation of transmitter release; a3b2 subtype composition has been suggested for the hippocampal noradrenaline release (Vizi et al., 1995; Sershen et al., 1997) and the striatal dopamine release (Kulak et al., 1997). b2 subunit‐containing nAChRs are involved in the regulation of GABA release in the thalamus (Lena and Changeux, 1997; Lena et al., 1993). In the superior cervical ganglion, activation of nAChRs enhanced the electrical field stimulation‐ induced release of ACh most likely via a3a7b2 nAChRs (Liang and Vizi, 1997). Activation of nAChRs could induce an increase in intracellular Ca2þ level of presynaptic neurits via a7 nAChRs (McGehee et al., 1995; Gray et al., 1996). Presynaptic nAChRs may combine their actions with an effect on the uptake, which results in a compound release mechanism (Szasz et al., 2005). This could occur because of the relatively high Ca2þ entry compared with Naþ influx that shifts the Ca2þ/Naþ exchanger to pump out Ca2þ leading to an increased Naþ entry and an opposite drive of the transporter. We may assume that if the Naþ concentration is high enough around the transporter it may change the direction of transport to release the transmitter instead of the uptake.

6.6 Postsynaptic Nicotinic Receptors There is evidence that a7‐ and b2‐containing nAChRs exist in both synaptic and nonsynaptic localizations in hippocampal neurons (Hill et al., 1993; Frazier et al., 1998b; Adams et al., 2001; Fabian‐Fine et al., 2001; Kawai et al., 2002; Graham et al., 2003). Most synapses containing nAChRs are not cholinergic but rather

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belong to glutamatergic or GABAergic presynaptic partners (Fabian‐Fine et al., 2001). For these receptors, ACh must diffuse in the synaptic cleft from the extracellular space. Thus the transmission in which they are involved can be taken as a specific form of nonsynaptic communication, nonsynaptic modulation of the postsynaptic area. Besides the studies on the hippocampus, there are a number of observations for the existence of nAChRs at nonsynaptic sites (Hill et al., 1993; Horch and Sargent, 1995; Ullian and Sargent, 1995; Williams et al., 1998; Fabian‐Fine et al., 2001). In chicken, ciliary ganglion nAChRs contribute to both synaptic and nonsynaptic transmission; a3/a5 nAChR mediate synaptic responses, while a7 nAChRs appear in perisynaptic locations (Horch and Sargent, 1995; Williams et al., 1998). Most nAChRs, which receive ACh message from presynaptic cholinergic boutons, are not true ‘‘postsynaptic’’ receptors, but are mostly located on extrasynaptic membranes. Given the nonsynaptic nature of nicotinic transmission in the CNS, it is more precise to define them as somatodendritic receptors. Hippocampal GABAergic inhibitory interneurons can be excited by activation of nAChRs (Alkondon et al., 1997; Jones and Yakel, 1997; Frazier et al., 1998b; McQuiston and Madison, 1999). These stratum radiatum interneurons likely mediate feed‐forward inhibition primarily because they receive inputs from fibers entering the CA1 and inhibit mostly dendrites of pyramidal neurons. There is heterogeneity in the types of interneurons regarding the nicotine sensitivity; all interneurons in the stratum radiatum and the stratum lacunosum moleculare can be excited by nicotinic ligands but many interneurons in the pyramidal cell layer do not respond to nicotine (McQuiston and Madison, 1999). Functional dendritic nAChRs exist in pyramidal neurons of the hippocampus and cause excitatory actions (Ji et al., 2001; Ge and Dani, 2005). Nicotinic excitation in interneurons can be amplified by other factors; simultaneous activation of AMPA and NMDA receptors boosts the postsynaptic nicotinic current in interneurons of the hippocampus (Alkondon et al., 2003). Not only are hippocampal interneurons highly sensitive to nicotinic stimulation; different types of cortical layer 5 interneurons can be excited by stimulation of nAChRs (Xiang et al., 1998).

6.7 Role of Nicotinic Receptors in Reward Mechanisms Tobacco use is driven by the rewarding effects of nicotine in the brain. Nicotine is an addictive drug that reinforces self‐administration and increases locomotion. Nicotine elevates the level of dopamine in the nucleus accumbens (NAc), which in turn reinforces the drug use particularly during the acquisition phase (Dani et al., 2001). Chronic exposure to nicotine desensitizes nAChRs, and over the long term, nAChRs enter long‐lasting inactive states (Pidoplichko et al., 1997). Meanwhile, an increase in number of nAChRs can be observed with the possible purpose to maintain the level of excitability (Dani and Heinemann, 1996). Nicotine‐induced limbic dopamine release may drive tobacco use, while inactivation of nAChRs due to the sustained low‐dose nicotine plays a role in tolerance and withdrawal. The addictive power of nicotine during smoking may be linked to the known potential of nAChRs to improve synaptic plasticity in regions of the brain reward system such as the VTA and the NAc. Blood level of nicotine in smokers, ranging between 250 and 500 nM for about 10 min just after smoking a cigarette (Henningfield et al., 1993), may reach the firing threshold of mesolimbic dopaminergic neurons leading to dopamine release in the nucleus accumbens. The role of b2 nAChR is well established in the neural mechanisms of reward (Picciotto et al., 1998). Recovery of the endogenous cholinergic transmission may require 30 min following the elimination of nicotine.

6.8 Special Role of Nicotinic Receptors in Neural Plasticity The simplest case of cellular nicotinic function related to plasticity is the well‐described enhancement in the frequency of synaptic activity that outlasts for several minutes after the end of the stimulation (Radcliffe and Dani, 1998; Sharma and Vijayaraghavan, 2003). Nicotine can also enhance the short‐term depression observed at thalamocortical connections (Gil et al., 1997). Presynaptic a7 nAChRs are able to induce long‐ term potentiation in the mesolimbic system by pairing nicotine application to postsynaptic depolarization of dopaminergic neurons (Mansvelder and McGehee, 2000). Nicotinic stimulation can also induce transition of short‐term potentiation of synaptic potentials into long term in the hippocampus (Ji et al., 2001;

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Mann and Greenfield, 2003). nAChRs use another possible mechanism to induce plastic changes in the hippocampus; nAChRs can transform the so‐called ‘‘silent’’ glutamatergic synapses into active synapses in CA1 pyramidal neurons (Maggi et al., 2003).

6.9 Nicotinic Receptors in Synaptic and Nonsynaptic Transmission There are experimental evidences for functional nAChRs, which mediate synaptic transmission. Cell types exhibiting nicotinic synaptic current include stratum radiatum and oriens interneurons (Frazier et al., 1998a), hippocampal CA1 pyramidal neurons (Hefft et al., 1999), pyramidal neurons and interneurons of the visual cortex (Roerig et al., 1997), and cells in the supraoptic nucleus (Hatton and Yang, 2002) and in the ciliary ganglion (Shoop et al., 2001). Both a7 and non‐a7 nAChRs can occur in synaptic transmission; in chick ciliary ganglion neurons, which innervate the iris and the choroid body, perisynaptic a7 nAChRs can also contribute to the fast synaptic current mediated by non‐a7 nAChRs (Chang and Berg, 1999). The dominant mode of the nicotinic influence on synaptic transmission, especially in the presynaptic actions, which, by nature, cannot be synaptic, seems to be the nonsynaptic information exchange. Activation of a7 nAChRs facilitates glutamatergic synaptic currents via a presynaptic action mechanism in cultured hippocampal neurons (Radcliffe and Dani, 1998), in CA1 and CA3 pyramidal neurons of the hippocampus (Ji et al., 2001; Maggi et al., 2003; Sharma and Vijayaraghavan, 2003), in pyramidal neurons of the rat auditory cortex (Aramakis and Metherate, 1998), and in mesolimbic dopaminergic neurons (Mansvelder and McGehee, 2000). Ca2þ influx seems to be a key player in the presynaptic mechanism of nAChR‐evoked responses. Low‐dose nicotine can increase frequency of miniature synaptic events, which are attributable to presynaptic activity, parallel with an increase in the presynaptic Ca2þ influx (McGehee et al., 1995; Gray et al., 1996; Maggi et al., 2003). There could be postsynaptic amplification by nAChRs, as well; low‐dose nicotine may enhance synaptic transmission by increasing the amplitude of evoked glutamatergic EPSCs via postsynaptic nAChRs in the interpeduncular nucleus (McGehee et al., 1995).

7

Muscarinic ACh Receptors

7.1 Subunits and Subtypes mAChRs received their name by their ability to bind muscarine, a product of the mushroom Amanita muscarina. mAChRs associate to G proteins and consist of five different receptors (M1–M5). Similar to nAChRs, a single neuron can express more than one mAChR subtype; for example, hippocampal pyramidal neurons have all five types of mAChRs (Levey et al., 1995). M1 receptors were first inactivated by genetic manipulation (Hamilton et al., 1997). Experiments on the M1 knockout mice revealed that seizure activity by muscarinic stimulation mostly connected to M1 receptors. In the striatum, M1 receptors might play a role in the early stages of Parkinson’s disease by increasing the dopamine release (Wess, 2003). Long‐ term potentiation in hippocampus is also reduced in M1 knockouts that exhibit a mild cognitive deficit in behavioral tests, suggesting the role of M1 receptors in learning and memory‐related processes (Anagnostaras et al., 2003). M2 receptor function has been associated with muscarinic stimulation‐induced tremor and akinesia. The block of VSCCs by muscarinic stimulation seems to be connected to activation of M2 receptors. M2 receptors participate in the presynaptic inhibition of transmitter release including ACh release. These autoreceptors of cholinergic transmission mediate feedback inhibition in the nervous system and in the neuromuscular junction. M3 receptors are widely expressed in the brain; the expression level is lower compared with other mAChR subtypes though. Adult M3 receptor‐deficient mice exhibit a weight loss of
25% (Yamada et al., 2001a). M3 receptors seem to play an important role in the regulation of appetite and daily food intake.

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M3 receptors also mediate the contractile response of smooth muscle by cholinergic stimulation in various areas including urinary bladder, ileum, and trachea, and are involved in parasympathetic control of pupillary sphincter muscle contractility in the eye (Wess, 2003). M4 receptors, similarly to M2 receptors, participate in the autoinhibition of cholinergic axons. These receptors also occur as presynaptic heteroreceptors, for example, in the regulation of dopamine release. M4 receptors influence locomotion through an interaction with D1 dopamine receptors. M5 receptors were the last muscarinic receptors to be cloned; it is not surprising that the physiological role of this receptor is less understood. In addition, their low expression level in the brain and the lack of specific ligand make them difficult to study. M5 receptors mediate ACh‐induced dilation of cerebral blood vessels (Yamada et al., 2001b) that might be important in the development of Alzheimer’s disease. M5 receptor is the sole mAChR subtype expressed by the substantia nigra dopaminergic neurons. This observation is particularly important as the loss of the nigrostriatal dopaminergic input is the cellular defect underlying Parkinson’s disease and ligands targeting mAChRs are used in the therapy of this disease. M5 receptors are involved in the oxotremorine‐induced facilitation of dopamine release in the striatum (Yamada et al., 2001b). Nevertheless, indirect effects through M4 receptors on GABAergic cells also contribute to the oxotremorine‐induced facilitation.

7.2 Subcellular Action Mechanisms M1, M3, and M5 receptors couple to Gq proteins and their activation mobilizes intracellular Ca2þ through a store‐mediated release. Through Gq/11, these mAChRs activate phospholipase C, which initiates the phosphatidylinositol turnover and produces inositol trisphosphate (IP3)‐mediated Ca2þ release from the intracellular Ca2þ stores, such as the endoplasmic reticulum. The breakdown of phosphatidylinositol also leads to diacylglycerol production, which activates protein kinase C and initiates a number of downstream cellular effectors. Overall, M1 or M3 receptors can induce ‘‘slow’’ increase in neuronal excitability by the higher Ca2þ level. M2 and M4 mAChRs can activate Kþ channels of the plasma membrane through Gi proteins resulting in hyperpolarization that can be seen as inhibitory effects on neural activity. As a consequence, M2 and M4 receptors inhibit action potential firing. One or both of these subtypes are found presynaptically on cholinergic (autoreceptors) and other transmitter‐containing (heteroreceptors) axon terminals where they inhibit neurotransmitter release. In the subcellular level, activation of these mAChRs causes activation of inward rectifying Kþ channels, inhibition of Ca2þ channels, and inhibition of adenylate cyclase and the subsequent intracellular processes. Not only direct potentiation of Kþ channels can be used to hyperpolarize target cells but the increases in intracellular free Ca2þ by M1 receptors can also activate Kþ current. M3 mAChRs are able to form functional dimers that make the mAChR‐mediated function more complex in vivo (Wess, 2003). M1 and M2 mAChRs in neurons and M2 and M3 mAChRs in smooth muscle cells may also heterodimerize. The functional role of dimerization of mAChRs are not known. Although ACh is rapidly hydrolyzed after release, desensitization of mAChRs occurs under physiological conditions. As with a large number of G‐protein coupled receptors, agonist‐induced desensitization of mAChRs usually involves receptor phosphorylation (Haga and Haga, 1990). The M1 and M3 mAChRs have been shown to be phosphorylated by protein kinase C (PKC). This receptor modification occurs on serine and threonine residues in the third cytoplasmic loop and the C‐terminus of the mAChRs. An array of protein kinases is able to phosphorylate mAChRs, including various G‐protein coupled receptor kinases (GRKs), casein kinase 1a (CK1a), and diacylglycerol‐regulated PKC. The mitogen‐activated protein kinases ERK1/2 phosphorylate a serine residue in GRK2, which decreases the activity of the kinase toward G‐protein coupled receptor substrates. Cytosolic b‐arrestin interacts with the phosphorylated receptor, leading to uncoupling of the mAChRs from the G proteins and clathrin‐coated vesicle formation. Following release of dynamin, clathrin, and b‐arrestin, the vesicle recycles back to the plasma membrane. Receptor internalization may represent a molecular mechanism appropriate for selective attenuation of particular mAChR signaling pathways.

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7.3 Pharmacology of Muscarinic Receptors General muscarinic agonists include several alkaloids, such as muscarine, arecoline, and synthetic compounds, such as carbachol. Bethanechol, metoclopramide, pilocarpine, and oxotremorine are other frequently used nonselective agonists of mAChRs (M1–M5). Absolute selectivity for any muscarinic agent has so far not been achieved. Subtype‐selective agonists are relatively rare; McN‐A‐343 is selective for the M1 receptor. There is more possibility to identify muscarinic subtypes with the use of selective antagonists. Subtypes of mAChRs can be identified by pharmacological tools (> Table 5-1): M1 receptors exhibit selectivity to the antagonist pirenzepine, M2 receptors can be selectively blocked by AF‐DX 116, M3 receptors can be selectively blocked by 4‐DAMP, whereas himbacine shows high affinity for the M4 receptor. M5 receptor has no selective ligand currently, and participation of this subtype is assumed when mAChR‐ mediated action is likely but all of the listed subtype‐selective antagonists are ineffective. The most used nonselective antagonists for mAChRs are atropine and scopolamine. Atropine, scopolamine, pirenzepine, and pilocarpine bind on the same binding site as the agonist but produce different conformational changes in the receptor structure that ultimately leads to the inhibition of the receptor function (van Koppen and Kaiser, 2003). All mAChR subtypes are susceptible to allosteric modulation; the binding of the allosteric modulator to the allosteric binding site results in a change in the conformation of the classical binding site leading to a change in the affinity of the receptor for classical muscarinic agonists and antagonists. There are also allosteric agents with positive cooperation on the binding of mAChR antagonists, such as strychnine (Lazareno and Birdsall, 1995). Other compounds, such as brucine, vincamine, and alcuronium, are able to allosterically modify the binding of mAChR agonists (Jakubik et al., 1997).

7.4 Muscarinic Receptor Functions mAChRs are responsible for postganglionic parasympathetic neurotransmission. Some sympathetic responses, such as sweating and piloerection, are also mediated by mAChRs. Although the ganglionic transmission is mediated by nAChRs by producing fast excitatory synaptic potential (EPSP) on the postsynaptic neuron, there is also a slow EPSP mediated by mAChRs. The slow EPSP decays in 1 sec and has a duration of 30–60 sec. Muscarinic agonists, acting on these ganglionic mAChRs, can enhance fast EPSPs under conditions of repetitive stimulation. It is important to note that not all of the peripherial organs are equipped with receptors to both sympathetic and parasympathetic transmitters. Therefore, nicotine‐sensitive ganglionic stimulation may end up with a non‐cholinergic effect, for example, in the blood vessels. M3 and M2 mAChRs are enriched in airway smooth muscle. In the CNS, the presynaptic mAChR (M2) inhibits cholinergic boutons as a muscarinic autoreceptor. Antagonism of this receptor by scopolamine results in release of choline into the extracellular space (Sarter and Parikh, 2005). It is well known that mAChRs play an important role in spike frequency adaptation in central neurons (Nicoll et al., 1990). Galantamine, a third generation cholinesterase inhibitor used in the therapy of Alzheimer’s disease, could dose dependently reduce the after hyperpolarization after a burst of action potentials and the spike‐ frequency accommodation of hippocampal CA1 neurons (Oh et al., 2005). Larger trains of backpropagating action potentials, exhibiting adaptation, were shown to be subject to modulation by mAChRs suggesting that dendritic integration can be modified by mAChRs (Tsubokawa and Ross, 1997). The cholinesterase blocker physostigmine and cholinomimetics evokes theta wave activity in the hippocampus through a muscarinic mechanism (Olpe et al., 1987; Konopacki et al., 1988; Golebiewski et al., 2002; Yoder and Pang, 2005). The rhythm is believed to be critical for the temporal coding or decoding of active neuronal ensembles and the modification of synaptic weights. Muscarinic antagonists, such as scopolamine or atropine, impair cognitive abilities in humans (Drachman, 1977). Recently it has been shown that transient activation of M1 mAChRs induces Ca2þ release from intracellular stores via IP3 and subsequent activation of an SK‐type Ca2þ‐activated Kþ conductance showing that ACh can directly inhibit neocortical pyramidal neurons through Ca2þ mobilization (Gulledge and Stuart, 2005). In the sensory system, presence of mAChRs has been shown on vestibular hair cells to evoke transmitter release from these cells (Derbenev et al., 2005).

Gene G protein Subcellular effect Selective agonist Selective antagonists

McN‐A‐343 Pilocarpine L‐689,660 Xanomeline Pirenzepine Telenzepine

M1 M1 Gq/11 Ic. Ca2þ (þ)

AF‐DX 116 Methoctramine AF‐DX 384 Gallamine Himbacine Tripitramine

M2 m2 Gi Kþ channels (þ) Ca2þ channels () adnylate cyclase () Bethanechol

. Table 5-1 mAChRs: basic pharmacology and subcellular effects of different subtypes

–

Himbacine Tropicamide AF‐DX 384

4‐DAMP Hexahydro‐sila‐ difenidol

L‐689,660

M5 m5 Gq/11 Ic. Ca2þ (þ) –

M4 m4 Gi Kþ channels (þ) Ca2þ channels () adnylate cyclase () McN‐A‐343

M3 m3 Gq/11 Ic. Ca2þ (þ)

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Conclusions

There are several examples that the effectors of the cholinergic, transmission, namely the muscarinic, and the nicotinic systems cooperate in the CNS. Although mAChRs are metabotropic receptors and mediate slow responses with considerable delay, nicotinic facilitation, following fast activation of nAChRs, can be sustained for up to 2 h. Interestingly, muscarinic inhibition seems to be more transient in certain cells (Girod and Role, 2001). Nicotinic agonists depolarize striatal interneurons and induce firing through non‐a 7 nAChRs, which, together with presynaptic inhibition through muscarinic receptors, form a dual cholinergic control on spiny interneurons of the striatum (Koos and Tepper, 2002). Main threads of current theories of nicotinic functions in the CNS include the following: (1) nAChRs may modulate, rather than mediate, fast synaptic transmission (McGehee and Role, 1995), (2) desensitization of the nAChRs, i.e., loss of function, is a key factor in the effect of nicotine during smoking and also shape the nAChR‐mediated activity in normal cholinergic transmission extending the computational power of the neuron, and (3) nAChRs directly release transmitters from presynaptic boutons skipping postsynaptic secondary modulations (Vizi and Lendvai, 1999). Taking these data together, we can conclude that the basis of the well‐known nicotinic enhancement of memory and learning function (Levin and Rezvani, 2000) is constructed on the level of cellular synaptic plasticity. Nicotine selectively improves the cognitive performance (especially those involving attentional processes) in deprived smokers and in cases with impaired cognition (Freedman et al., 1995), such as Alzheimer’s disease (Rezvani and Levin, 2001; Newhouse et al., 2004). Cholinergic therapy is the mainstay treatment for Alzheimer’s disease. In conclusion, cholinergic system plays an effective role in modulating cognitive functions in the brain acting on a rich assembly of metabotropic and ionotropic receptors. The current view that nonsynaptic communication is dominant in cholinergic transmission also supports the future perspective of drug therapy targeting high‐affinity nonsynaptic receptors.

References Adams CE, Stitzel JA, Collins AC, Freedman R. 2001. Alpha7‐ nicotinic receptor expression and the anatomical organization of hippocampal interneurons. Brain Res 922: 180-190. Alkondon M, Pereira EF, Albuquerque EX. 2003. NMDA and AMPA receptors contribute to the nicotinic cholinergic excitation of CA1 interneurons in the rat hippocampus. J Neurophysiol 90: 1613-1625. Alkondon M, Pereira EF, Cortes WS, Maelicke A, Albuquerque EX. 1997. Choline is a selective agonist of a7 nicotinic acetylcholine receptors in the rat brain neurons. Eur J Neurosci 9: 2734-2742. Anagnostaras SG, Murphy GG, Hamilton SE, Mitchell SL, Rahnama NP, et al. 2003. Selective cognitive dysfunction in acetylcholine M1 muscarinic receptor mutant mice. Nat Neurosci 6: 51-58. Aramakis VB, Metherate R. 1998. Nicotine selectively enhances NMDA receptor‐mediated synaptic transmission during postnatal development in sensory neocortex. J Neurosci 18: 8485-8495. Bennett BD, Wilson CJ. 1999. Spontaneous activity of neostriatal cholinergic interneurons in vitro. J Neurosci 19: 5586-5596. Chang KT, Berg DK. 1999. Nicotinic acetylcholine receptors containing a7 subunits are required for reliable synaptic transmission in situ. J Neurosci 19: 3701-3710.

Chiodini FC, Tassonyi E, Hulo S, Bertrand D, Muller D. 1999. Modulation of synaptic transmission by nicotine and nicotinic antagonists in hippocampus. Brain Res Bull 48: 623-628. Dani JA, Heinemann S. 1996. Molecular and cellular aspects of nicotine abuse. Neuron 16: 905-908. Dani JA, Ji D, Zhou FM. 2001. Synaptic plasticity and nicotine addiction. Neuron 31: 349-352. Derbenev AV, Linn CL, Guth PS. 2005. Muscarinic ACh receptor activation causes transmitter release from isolated frog vestibular hair cells. J Neurophysiol 94: 3134-3142. Descarries L, Mechawar N. 2000. Ultrastructural evidence for diffuse transmission by monoamine and acetylcholine neurons of the central nervous system. Prog Brain Res 125: 27-47. Didier M, Berman SA, Lindstrom J, Bursztajn S. 1995. Characterization of nicotinic acetylcholine receptors expressed in primary cultures of cerebellar granule cells. Mol Brain Res 30: 17-28. Dong H, Xiang YY, Farchi N, Ju W, Wu Y, et al. 2004. Excessive expression of acetylcholinesterase impairs glutamatergic synaptogenesis in hippocampal neurons. J Neurosci 24: 8950-8960. Drachman DA. 1977. Memory and cognitive function in man: Does the cholinergic system have a specific role? Neurology 27: 783-790.

Cholinergic transmission Eckenstein F, Baughman RW. 1984. Two types of cholinergic innervation in cortex, one co‐localized with vasoactive intestinal polypeptide. Nature 309: 153-155. Fabian‐Fine R, Skehel P, Errington ML, Davies HA, Sher E, et al. 2001. Ultrastructural distribution of the a7 nicotinic acetylcholine receptor subunit in rat hippocampus. J Neurosci 21: 7993-8003. Frazier CJ, Buhler AV, Weiner JL, Dunwiddie TV. 1998a. Synaptic potentials mediated via a‐bungarotoxin‐sensitive nicotinic acetylcholine receptors in rat hippocampal interneurons. J Neurosci 18: 8228-8235. Frazier CJ, Rollins YD, Breese CR, Leonard S, Freedman R, et al. 1998b. Acetylcholine activates an a‐bungarotoxin‐ sensitive nicotinic current in rat hippocampal interneurons, but not pyramidal cells. J Neurosci 18: 1187-1195. Freedman R, Hall M, Adler LE, Leonard S. 1995. Evidence in postmortem brain tissue for decreased numbers of hippocampal nicotinic receptors in schizophrenia. Biol Psychiatry 38: 22-33. Ge S, Dani JA. 2005. Nicotinic acetylcholine receptors at glutamate synapses facilitate long‐term depression or potentiation. J Neurosci 25: 6084-6091. Gil Z, Connors BW, Amitai Y. 1997. Differential regulation of neocortical synapses by neuromodulators and activity. Neuron 19: 679-686. Girod R, Role LW. 2001. Long‐lasting enhancement of glutamatergic synaptic transmission by acetylcholine contrasts with response adaptation after exposure to low‐level nicotine. J Neurosci 21: 5182-5190. Golebiewski H, Eckersdorf B, Konopacki J. 2002. Septal cholinergic mediation of hippocampal theta in the cat. Brain Res Bull 58: 323-335. Graham AJ, Ray MA, Perry EK, Jaros E, Perry RH, et al. 2003. Differential nicotinic acetylcholine receptor subunit expression in the human hippocampus. J Chem Neuroanat 25: 97-113. Gray R, Rajan AS, Radcliffe KA, Yakehiro M, Dani JA. 1996. Hippocampal synaptic transmission enhanced by low concentrations of nicotine. Nature 383: 713-716. Gulledge AT, Stuart GJ. 2005. Cholinergic inhibition of neocortical pyramidal neurons. J Neurosci 25: 10308-10320. Haga K, Haga T. 1990. Dual regulation by G proteins of agonistdependent phosphorylation of muscarinic acetylcholine receptors. FEBS Lett 268: 43-47. Hamilton SE, Loose MD, Qi M, Levey AI, Hille B, et al. 1997. Disruption of the m1 receptor gene ablates muscarinic receptor‐dependent M current regulation and seizure activity in mice. Proc Natl Acad Sci USA 94: 13311-13316. Hatton GI, Yang QZ. 2002. Synaptic potentials mediated by a7 nicotinic acetylcholine receptors in supraoptic nucleus. J Neurosci 22: 29-37.

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Hefft S, Hulo S, Bertrand D, Muller D. 1999. Synaptic transmission at nicotinic acetylcholine receptors in rat hippocampal organotypic cultures and slices. J Physiol 515: 769-776. Henningfield JE, Stapleton JM, Benowitz NL, Grayson RF, London ED. 1993. Higher levels of nicotine in arterial than in venous blood after cigarette smoking. Drug Alcohol Depend 33: 23-29. Hill JA, Zoli M, Bourgeois J‐P, Changeux J‐P. 1993. Immunocytochemical localization of neural nicotinic receptor: The b2‐subunit. J Neurosci 13: 1551-1568. Horch HL, Sargent PB. 1995. Perisynaptic surface distribution of multiple classes of nicotinic acetylcholine receptors on neurons in the chicken ciliary ganglion. J Neurosci 15: 7778-7795. Jakubik J, Bacakova L, El‐Fakahany EE, Tucek S. 1997. Positive cooperativity of acetylcholine and other agonists with allosteric ligands on muscarinic acetylcholine receptors. Mol Pharmacol 52: 172-179. Ji D, Lape R, Dani JA. 2001. Timing and location of nicotinic activity enhances or depresses hippocampal synaptic plasticity. Neuron 31: 131-141. Jones S, Yakel JL. 1997. Functional nicotinic ACh receptors on interneurones in the rat hippocampus. J Physiol 504: 603-610. Kawai H, Zago W, Berg DK. 2002. Nicotinic a7 receptor clusters on hippocampal GABAergic neurons: Regulation by synaptic activity and neurotrophins. J Neurosci 22: 7903-7912. Khiroug L, Giniatullin R, Klein RC, Fayuk D, Yakel JL. 2003. Functional mapping and Ca2þ regulation of nicotinic acetylcholine receptor channels in rat hippocampal CA1 neurons. J Neurosci 23: 9024-9031. Khiroug L, Giniatullin R, Sokolova E, Talantova M, Nistri A. 1997. Imaging of intracellular calcium during desensitization of nicotinic acetylcholine receptors of rat chromaffin cells. Br J Pharmacol 122: 1323-1332. Kiss JP, Vizi ES, Westerink BHC. 1999. Effect of neostigmine on the hippocampal noradrenaline release: Role of cholinergic receptors. Neuroreport 10: 81-86. Kofalvi A, Sperlagh B, Zelles T, Vizi ES. 2000. Long‐lasting facilitation of 4‐amino‐n‐[2,3‐3H]butyric acid ([3H] GABA) release from rat hippocampal slices by nicotinic receptor activation. J Pharmacol Exp Ther 295: 453-462. Konopacki J, Bland BH, Roth SH. 1988. Evidence that activation of in vitro hippocampal theta rhythm only involves muscarinic receptors. Brain Res 455: 110-114. Koos T, Tepper JM. 2002. Dual cholinergic control of fast‐ spiking interneurons in the neostriatum. J Neurosci 22: 529-535. Kulak JM, Nguyen TA, Olivera BM, McIntosh JM. 1997. a‐ Conotoxin MII blocks nicotine‐ stimulated dopamine release in rat striatal synaptosomes. J Neurosci 17: 5263-5270.

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Lazareno S, Birdsall NJ. 1995. Detection, quantitation, and verification of allosteric interactions of agents with labeled and unlabeled ligands at G protein‐coupled receptors: Interactions of strychnine and acetylcholine at muscarinic receptors. Mol Pharmacol 48: 362-78. Lena C, Changeux J‐P, Mulle C. 1993. Evidence for ‘preterminal’ nicotine receptors on GABAergic axons in the rat interpeduncular nucleus. J Neurosci 13: 2680-2688. Lena C, Changeux J-P. 1997. Role of Ca2+ ions in nicotinic facilitation of GABA release in mouse thalamus. J Neurosci 17: 576-585. Lendvai B, Sershen H, Lajtha A, Santha E, Baranyi M, et al. 1996. Differential mechanisms involved in the effect of nicotinic agonists DMPP and lobeline to release [3H]5‐ HT from rat hippocampal slices. Neuropharmacology 35: 1769-1777. Lester RA, Dani JA. 1994. Time‐dependent changes in central nicotinic acetylcholine channel kinetics in excised patches. Neuropharmacology 33: 27-34. Levey AI, Edmunds SM, Koliatsos V, Wiley RG, Heilman CJ. 1995. Expression of m1‐m4 muscarinic acetylcholine receptor proteins in rat hippocampus and regulation by cholinergic innervation. J Neurosci 15: 4077-4092. Levin ED, Rezvani AH. 2000. Development of nicotinic drug therapy for cognitive disorders. Eur J Pharmacol 393: 141-146. Liang SD, Vizi ES. 1997. Positive feedback modulation of acetylcholine release from isolated superior cervical ganglion. J Pharmacol Exp Ther 280: 650-655. Lukas RJ. 1991. Effects of chronic nicotinic ligand exposure on functional activity of nicotinic acetylcholine receptors expressed by cells of the PC12 rat pheochromocytoma or the TE671/RD human clonal line. J Neurochem 56: 1134-1145. Maggi L, Le Magueresse C, Changeux JP, Cherubini E. 2003. Nicotine activates immature ‘‘silent’’ connections in the developing hippocampus. Proc Natl Acad Sci USA 100: 2059-2064. Mann EO, Greenfield SA. 2003. Novel modulatory mechanisms revealed by the sustained application of nicotine in the guinea‐pig hippocampus in vitro. J Physiol 551.2: 539-550. Mansvelder HD, McGehee DS. 2000. Long‐term potentiation of excitatory inputs to brain reward areas by nicotine. Neuron 27: 349-357. McGehee DS, Role LW. 1995. Physiological diversity of nicotinic acetylcholine receptors expressed by vertebrate neurons. Annu Rev Physiol 57: 521-546. McGehee DS, Heath MJ, Gelber S, Devay P, Role LW. 1995. Nicotine enhancement of fast excitatory synaptic transmission in CNS by presynaptic receptors. Science 269: 1692-1696.

McQuiston AR, Madison DV. 1999. Nicotinic receptor activation excites distinct subtypes of interneurons in the rat hippocampus. J Neurosci 19: 2887-2896. Mulle C, Choquet D, Korn H, Changeux JP. 1992. Calcium influx through nicotinic receptor in rat central neurons: Its relevance to cellular regulation. Neuron 8: 135-143. Newhouse PA, Potter A, Singh A. 2004. Effects of nicotinic stimulation on cognitive performance. Curr Opin Pharmacol 4: 36-46. Nicoll RA, Malenka RC, Kauer JA. 1990. Functional comparison of neurotransmitter receptor subtypes in mammalian central nervous system. Physiol Rev 70: 513-565. Oh MM, Wu WW, Power JM, Disterhoft JF. 2005. Galantamine increases excitability of CA1 hippocampal pyramidal neurons. Neuroscience Oct 19; [Epub ahead of print]. Olpe HR, Klebs K, Kung E, Campiche P, Glatt A, et al. 1987. Cholinomimetics induce theta rhythm and reduce hippocampal pyramidal cell excitability. Eur J Pharmacol 142: 275-283. Picciotto MR, Zoli M, Rimondini R, Lena C, Marubio LM, et al. 1998. Acetylcholine receptors containing the b2 subunit are involved in the reinforcing properties of nicotine. Nature 391: 173-177. Pidoplichko VI, De Biasi M, Williams JT, Dani JA. 1997. Nicotine activates and desensitizes midbrain dopamine neurons. Nature 390: 401-404. Radcliffe KA, Dani JA. 1998. Nicotinic stimulation produces multiple forms of increased glutamatergic synaptic transmission. J Neurosci 18: 7075-7083. Rezvani AH, Levin ED. 2001. Cognitive effects of nicotine. Biol Psychiatry 49: 258-267. Roerig B, Nelson DA, Katz LC. 1997. Fast synaptic signaling by nicotinic acetylcholine and serotonin 5‐HT3 receptors in developing visual cortex. J Neurosci 17: 8353-8362. Role LW, Berg DK. 1996. Nicotinic receptors in the development and modulation of CNS synapses. Neuron 16: 1077-1085. Rowell PP, Duggan DS. 1998. Long‐lasting inactivation of nicotinic receptor function in vitro by treatment with high concentrations of nicotine. Neuropharmacology 37: 103-111. Sarter M, Parikh V. 2005. Choline transporters, cholinergic transmission and cognition. Nat Rev Neurosci 6: 48-56. Seguela P, Wadiche J, Dineley‐Miller K, Patrick JW. 1993. Molecular cloning, functional properties, and distribution of rat brain a7: A nicotinic cation channel highly permeable to calcium. J Neurosci 13: 596-604. Sershen H, Balla A, Lajtha A, Vizi ES. 1997. Characterization of nicotinic receptors involved in the release of norepinephrine from hippocampus. Neuroscience 77: 121-130.

Cholinergic transmission Sharma G, Vijayaraghavan S. 2003. Modulation of presynaptic store calcium induces release of glutamate and postsynaptic firing. Neuron 38: 929-939. Shoop RD, Chang KT, Ellisman MH, Berg DK. 2001. Synaptically driven calcium transients via nicotinic receptors on somatic spines. J Neurosci 21: 771-781. Soliakov L, Wonnacott S. 1996. Voltage sensitive Ca2þ channels involved in nicotinic receptor‐mediated 3Hdopamine release from rat striatal synaptosomes. J Neurochem 67: 163-170. Szasz BK, Mayer A, Zsilla G, Lendvai B, Vizi ES, et al. 2005. Carrier‐mediated release of monoamines induced by the nicotinic acetylcholine receptor agonist DMPP. Neuropharmacology 49: 400-409. Towart LA, Alves SE, Znamensky V, Hayashi S, McEwen BS, et al. 2003. Subcellular relationships between cholinergic terminals and estrogen receptor‐alpha in the dorsal hippocampus. J Comp Neurol 463: 390-401. Tsubokawa H, Ross WN. 1997. Muscarinic modulation of spike backpropagating in the apical dendrites of hippocampal CA1 pyramidal neurons. J Neurosci 17: 5782-5791. Ullian EM, Sargent PB. 1995. Pronounced cellular diversity and extrasynaptic location of nicotinic acetylcholine receptor subunit immunreactivities in the chicken pretectum. J Neurosci 15: 7012-7023. Umbriaco D, Garcia S, Beaulieu C, Descarries L. 1995. Relational features of acetylcholine, noradrenaline, serotonin and GABA axon terminals in the stratum radiatum of adult rat hippocampus (CA1). Hippocampus 5: 605-620. Umbriaco D, Watkins KC, Descarries L, Cozzari C, Hartman BK. 1994. Ultrastructural and morphometric features of the acetylcholine innervation in adult rat parietal cortex: An electron microscopic study in serial sections. J Comp Neurol 348: 351-373. van Koppen CJ, Kaiser B. 2003. Regulation of muscarinic acetylcholine receptor signaling. Pharmacol Ther 98: 197-220. Vernino S, Amador M, Luetje CW, Patrick J, Dani JA. 1992. Calcium modulation and high calcium permeability of neuronal nicotinic acetylcholine receptors. Neuron 8: 127-134.

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Vijayaraghavan S, Pugh PC, Zhang ZW, Rathouz MM, Berg DK. 1992. Nicotinic receptors that bind agr‐bungarotoxin on neurons raise intracellular free Ca2þ . Neuron 8: 353-362. Vizi ES. 2000. Role of high‐affinity receptors and membrane transporters in nonsynaptic communication and drug action in the central nervous system. Pharmacol Rev 52: 63-89. Vizi ES, Lendvai B. 1999. Modulatory role of presynaptic nicotinic receptors in synaptic and non‐synaptic chemical communication in the central nervous system. Brain Res Rev 30: 219-235. Vizi ES, Sershen H, Balla A, Mike A, Windish K, et al. 1995. Neurochemical evidence of heterogeneity of presynaptic and somatodendritic nicotinic acetylcholine receptors. Ann NY Acad Sci 757: 84-99. Wess J. 2003. Novel insights into muscarinic acetylcholine receptor function using gene targeting technology. Trends Pharmacol Sci 24: 414-420. Williams BM, Temburni MK, Levey MS, Bertrand S, Bertrand D, et al. 1998. The long internal loop of the a3 subunit targets nAChRs to subdomains within individual synapses on neurons in vivo. Nat Neurosci 1: 557-562. Wonnacott S. 1997. Presynaptic nicotinic ACh receptors. Trends Neurosci 20: 92-98. Xiang Z, Huguenard JR, Prince DA. 1998. Cholinergic switching within neocortical inhibitory networks. Science 281: 985-988. Yamada M, Lamping KG, Duttaroy A, Zhang W, Cui Y, et al. 2001b. Cholinergic dilation of cerebral blood vessels is abolished in M(5) muscarinic acetylcholine receptor knockout mice. Proc Natl Acad Sci USA 98: 14096-14101. Yamada M, Miyakawa T, Duttaroy A, Yamanaka A, Moriguchi T, et al. 2001a. Mice lacking the M3 muscarinic acetylcholine receptor are hypophagic and lean. Nature 410: 207-212. Yoder RM, Pang KC. 2005. Involvement of GABAergic and cholinergic medial septal neurons in hippocampal theta rhythm. Hippocampus 15: 381-392. Zhou FM, Liang Y, Dani JA. 2001. Endogenous nicotinic cholinergic activity regulates dopamine release in the striatum. Nat Neurosci 4: 1224-1229.

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Molecular Genetics of Brain Noradrenergic Neurotransmission

R. Meloni

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 130

2 2.1 2.2 2.3 2.4 2.5 2.6 2.6.1 2.6.2

Brain Noradrenergic System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 130 Ontogeny . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 130 Nuclei and Pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 131 NE Synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 131 NE Storage and Release . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 131 NE Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 132 Termination of NE Synaptic Transmission . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 133 Reuptake . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 133 Metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 134

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Functional Noradrenergic System Neurotransmission . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 134 Brain Noradrenergic Neurotransmission and Arousal . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 134 Tonic Versus Phasic Excitation of NE Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 134 Brain Noradrenergic Neurotransmission, Stress, and Depression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 135

4 4.1 4.2 4.3 4.4

Genetics of Noradrenergic Neurotransmission . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 135 Tyrosine Hydroxylase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 136 Dopamine b Hydroxylase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 137 Mono‐Amine‐Oxydase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 138 Catechol‐O‐Methyltransferase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 139

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Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 140

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Abstract: The brain noradrenergic system plays a pivotal role in mediating the responses of the organism to the external and internal milieu and particularly to stress. The different components of this system have been implicated in a wide range of normal and pathological behavior. Recent advances in molecular psychiatric genetics may pave the way for a better understanding of the etiology of different symptoms of mental diseases and their relationship to environmental factors. List of Abbreviations: AAD, Aromatic Amino acid Decarboxylase; AC, adenylyl cyclase; ADs, antidepressant drugs; BMP, Bone Morphogenetic Proteins; COMT, catechol‐O‐methyltransferase; DAG, diacylglycerol; DOPA, 3,4 dihydroxyphenylalanine; DHPG, 3,4‐dihydroxyphenyl‐glycol; MHPG, 3‐methoxy‐4‐ hydroxyphenylglycol; fmri, functional magnetic resonance imaging; Gata3, GATA‐binding protein 3; GDP, guanosine diphosphate; GPCRs, G protein‐coupled receptors; GTP, guanosine triphosphate; HPA, hypothalamic‐pituitary‐adrenal; LC, Locus Coeruleus; MAOs, monoamineoxidases; NET, NE transporter; PKC, protein kinase C; PKA, protein kinases A; PNMT, Phenylethanolamine‐N‐Methyl‐Transferase; PLC, phospholipase C; PTSD, posttraumatic stress disorder; SSRIs, selective serotonin reuptake inhibitors; TH, Tyrosine Hydroxylase; VMA, vanyl‐mandelic acid; VCF, velocardiofacial syndrome; VMAT, vesicular monoamine transporter

1

Introduction

Adrenaline (L(
)‐epinephrine) and noradrenaline (L(
)‐norepinephrine) (NE), along with dopamine, are catecholamines, a class of molecules characterized by a catechol and an amine group, which belong with the indolamines (such as serotonin) to the monoamines group of compounds. They act as neurotransmitters as well as hormones, NE being the predominant neurotransmitter whereas epinephrine is the major hormone. Their names derive from the ancient Greek ‘‘epi’’ (upon) and ‘‘nephron’’ (kidney), since epinephrine is produced mainly by the adrenal glands located in the apex of the kidneys. Epinephrine released from the adrenal glands is carried in the bloodstream and acts upon the autonomous nervous system regulating a wide range of functions such as the heart rate, dilation of the pupils, glucose storage, intestinal motility, secretion of sweat, and salivary glands etc. In the brain the cell bodies that contain NE are found in the brainstem, and their projections extend from the forebrain to the spinal cord. These neurons are associated with the stress response and with the control of drive and motivation, alertness and sleep patterns, along with stress‐related manifestations such as anxiety and fear. Taken together the physiological reactions in the peripheral and central nervous system mediated by the adrenergic and NE‐ergic systems make up the substrate of the response to stress that is best illustrated by the ‘‘Fight or Flight’’ paradigm. The central role of the stress response in the homeostasis of the living organism has placed it also at the crossroads of several peripheral and central pathologies that are totally or partially related to inadequate, excessive, or prolonged activation of the mechanism underlying stress. In the brain, stress has been associated with such pathologies as depression and anxiety, but also bipolar disorder and schizophrenia (Berridge and Waterhouse, 2003).

2

Brain Noradrenergic System

2.1 Ontogeny Mash‐1, a bHLH domain protein, as well as Phox2a and Phox2b, two paralogous homeodomain proteins belonging to the Q50 paired‐like class, are transcription factors essential for the development of all central and peripheral neurons with a NE‐ergic phenotype. These factors have been implicated in synchronizing pan‐neuronal and catecholaminergic specific neurogenesis. Their expression domain is restricted, in the peripheral nervous system, to all autonomic ganglia (sympathetic, parasympathetic, and enteric) and the distal ganglia of the facial, hypoglossal, and vagus cranial nerves. In the central nervous system, it comprises all NE‐ergic centers, all cranial motor nuclei with the exception of the abducens and hypoglossal,

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the nucleus of the solitary tract and associated area postrema, as well as scattered interneurons in the hindbrain and spinal cord (Brunet and Pattyn, 2002). The cascade of evenements leading to the NE‐ergic phenotype encompasses upstream Mash‐1 and Phox2a/2b, the bone morphogenetic proteins (BMP) class of factors and, downstream, the bHLH transcription factor dHand (heart and neural‐crest derivatives expressed 2), and the zinc finger protein GATA‐binding protein 3 (Gata3). In the periphery, Mash‐1 appears to regulate, with Phox2b, the expression of dHand and Phox2a. These two factors may interact and regulate with Gata3 the expression of the tyrosine hydroxylase (TH) and dopamine‐b‐hydroxylase genes, which constitute the hallmarks of the NE‐ergic phenotype. In the brain, this pathway appears to be more straightforward forming a direct chain with the factors acting in the sequence BMP, Mash‐1, Phox2a, and Phox2b. This latter then regulates (with Phox2a?) the expression of the NE‐ergic synthesizing enzyme tyrosine hydroxylase and dopamine‐b‐hydroxylase (Goridis and Rohrer, 2002). Interestingly, point mutations in the HASH‐1 gene and a polyalanine expansion in the PHOX2B gene product have been associated with congenital central hypoventilation syndrome in humans, a rare breathing disorder accompanied by dysautomia manifestations, which is also known as Ondine’s curse (de Pontual et al., 2003).

2.2 Nuclei and Pathways All NE‐ergic neurons in the brain are located in the brainstem. In the pons they form the locus coeruleus (LC), and in the medulla they are found in the reticular formation, the solitary nucleus, and the dorsal motor nucleus of the vagus. The NE‐ergic system originates in a relatively small number of cells, but it innervates, by generating an extensive network of NE‐ergic projections, essentially the whole neuraxis from the olfactory bulb to the spinal cord. In this way, this system may potentially influence the activity of a widespread array of brain centers under conditions of elevated NE‐ergic activity. The projections originating from the LC form the dorsal noradrenergic bundle, a pathway that merges with the median forebrain bundle, the medial forebrain bundle, and the dorsal longitudinal fasciculum. The LC is at the origin of most of the innervation to the forebrain, with over 40% of its neurons projecting to the neocortex (frontal lobes, hippocampus, and olfactory bulbs), representing the most important NE‐ergic input related to psychological functions (Aston‐Jones and Cohen, 2005). Other projections are directed to the basal forebrain, the thalamus, and the cerebellum, whereas another important output is toward the brainstem sensory and association nuclei and all the levels of the spinal cord, mediating autonomic regulation of, for example, the cardiac activity or the axis hypothalamic–pituitary–adrenal (HPA) stress axis. Afferents to the LC are inhibitory GABA‐ergic inputs from the rostral medulla and the nucleus prepositus hypoglossi, and excitatory inputs, probably glutamatergic, from the ventromedial zone of the nucleus paragigantocellularis.

2.3 NE Synthesis Tyrosine is the common precursor in the synthesis of NE and adrenaline, as well as dopamine. Tyrosine is converted inside the nerve terminal to 3,4‐dihydroxyphenylalanine (DOPA) by tyrosine hydroxylase (TH), the rate‐limiting enzyme in the synthesis of catecholamines. DOPA is then decarboxylated by the aromatic amino acid decarboxylase (AAD or DOPA decarboxylase) to dopamine, which is then converted to NE by dopamine‐b‐hydroxylase. Thereafter, the phenylethanolamine‐N‐methyl‐transferase (PNMT) converts the NE to adrenaline in the adrenal medulla.

2.4 NE Storage and Release NE, as all monoamines, is concentrated in vesicles at the nerve terminal by a specific vesicular monoamine transporter (VMAT‐1 and VMAT‐2) (Njus et al., 1986). VMAT‐1 is primarily present in endocrine and paracrine cells of peripheral organs, whereas VMAT‐2 is the predominant monoamine vesicular transporter in the central nervous system (Masson et al., 1999).

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The calcium inflow triggered by the opening of voltage‐activated calcium channels upon the arrival of an action potential to the nerve terminal induces the fusion of the vesicle with the presynaptic membrane and the release of the neurotransmitter into the synaptic cleft. There it will act on specific receptors located both on the postsynaptic and presynaptic membranes. Stimulation of the postsynaptic receptors results in changes in the properties of the postsynaptic membrane with either a shift in membrane potential when the receptors are coupled with ion channels (ionotropic receptors) or in biochemical changes when the receptors are coupled with G proteins (metabotropic receptors) (Starke, 2001). Stimulation of the presynaptic receptors located on the nerve terminal will regulate the transmitter release triggered by the action potential, providing, therefore, a feedback mechanism for the control of the neurotransmitter’s concentration in the synaptic cleft (Boehm and Kubista, 2002).

2.5 NE Receptors NE and adrenaline exert their cellular action via binding to specific membrane proteins, the adrenergic receptors or adrenoceptors. The adrenergic receptors are classified into two main categories, alpha and beta, and can be divided into three main classes based on sequence similarity, receptor pharmacology, and signaling mechanisms: alphal (a1), alpha2 (a2), and beta (b). All three classes are present in the brain. Further subdivisions exist within each class (Bylund, 1992; Bylund, 2005). All adrenergic receptors belong to the family of G protein‐coupled receptors (GPCRs), the largest single family of integral membrane receptors. It has been calculated that 3–4% of the genes encodes a member of this family. GPCRs are characterized by seven transmembrane domains and are coupled to heterotrimeric—since they are composed of three subunits (a, b, and g)—guanine nucleotide‐binding proteins (G proteins). The binding of the agonist to the receptor induces a conformational change characterized by a high affinity agonist state that turn G proteins on by promoting the binding of the activating nucleotide guanosine triphosphate (GTP) in exchange for guanosine diphosphate (GDP) on the a subunit of the G protein. This activation of the specific G protein initiates a cascade of molecular events resulting in the positive or negative regulation of the effectors systems (Neer, 1995). The G proteins are named after their a subunit and are divided into four subgroups: Gs and Gi/o, which stimulate and inhibit, respectively, adenylate cyclase, Gq/11, which stimulates phospholipase C (PLC), and the less characterized G12/13 subgroup, which activates the Naþ/Hþ exchanger pathway (Gether, 2000). The distinct classes of adrenergic receptors are differently coupled to these subgroups. The b‐adrenergic receptor, which presents three subtypes (b1, b2, and b3) and is exclusively postsynaptic, is coupled to the Gs proteins. The stimulation of the b‐adrenergic receptor leads to the prototypic cellular effect of Gs proteins, which is activation via stimulation of adenylyl cyclase (AC) resulting in the accumulation of the second messenger cyclic adenosine monophosphate (c‐AMP) and subsequent activation of the c‐AMP‐dependent protein kinases A (PKA). PKA causes the phosphorylation of various cellular proteins, which produce the specific responses of b‐adrenergic receptor stimulation (Taussig and Gilman, 1995). Molecular cloning has identified four different subtypes of a2 receptors designated as a2A, a2B, a2C, and a2D (Bylund et al., 1994). The a2 receptors are both presynaptic and postsynaptic and the a2A receptor appears to be far more represented in the brain than in the sympathetic nervous system (Boehm and Kubista, 2002). The a2 adrenergic receptors are coupled to the Gi/o protein family whose activation results in the inhibition of c‐AMP accumulation (Neer, 1995). The a2 adrenergic receptors mediate a hyperpolarization of the neuronal membrane, which renders the neuron less excitable by increasing Kþ conductance via activation of G protein‐gated Kþ channels. By their localization and action the presynaptic a2 receptors regulate the synthesis and release of NE. The al adrenergic receptors (comprising the 1A, 1B, 1C, and 1D subtypes) are generally excitatory in nature and are usually coupled to the Gi/Gq family of G proteins. The Gi/Gq proteins are linked to a signaling cascade different from the AC pathway. They activate the phosphatidylinositol‐specific PLC, which subsequently generates the second set of messengers, inositol triphosphate (IP3) and diacylglycerol (DAG). IP3 stimulates the release of Ca2þ from intracellular stores via a specific receptor‐mediated process,

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thus increasing available intracellular calcium that will be involved in the regulation of several protein kinases (Berridge, 1993). DAG is a potent activator of protein kinase C (PKC), which is involved in the activation of many substrates including membrane proteins such as channels, pumps, and ion exchange proteins (Fields and Casey, 1997). Stimulation of the membrane PLC leading to the formation of the second messengers IP3 and DAG through activation of the al receptors on central adrenergic target neurons produces depolarizing responses due to a decrease in Kþ conductance, thus making the neuron more excitable. The activation of a GPCR not only results in the G protein‐dependent activation of effector systems but also allows for feedback regulation of G protein coupling, receptor endocytosis, and signaling through G protein independent signal transduction pathways. Therefore, GPCR activity represents a coordinated balance between molecular mechanisms governing receptor signaling, desensitization, and resensitization as well as downregulation. Desensitization is the consequence of the uncoupling of the receptor from heterotrimeric G proteins in response to receptor phosphorylation by specific kinases (GRKs). This phosphorylation favorizes the interactions with G proteins to mediate the effects of receptor stimulation and, at the same time, target the receptor for the binding of the arrestin class of proteins. The arrestins bind to the phosphorylated receptor on the cytoplasmatic side and block further G protein activation, operating in this way to desensitize the receptor. Arrestins act subsequently as a molecular adaptor in binding with clathrin and the clathrin adaptor complex AP‐2. This produces the internalization by endocytosis of cell surface receptors and their sequestration in intracellular membranous compartments. Then the sequestered receptors can be recycled, producing resensitization, or directed to the lysosomial apparatus for degradation, resulting in a downregulation, which is a reduction of the total number of receptors. Conversely, after reduction in the chronic level of receptor stimulation by chronic antagonist administration or denervation, hyperreactivity and upregulation of the GPCRs are stimulated (Ferguson, 2001). The pivotal role played by arrestins in a NE‐ergic transmission, and in general in the activity of all GPCRs, is not limited just to the extinguishing of the signal. They have been shown to act also as a multifunctional adaptor and as scaffolding proteins implicated in recruiting signaling molecules. In mammals there are two types of arrestins: the visual arrestins that are mainly restricted to photoreceptor cells, and the b‐arrestins (1 and 2) that are ubiquitously expressed. Upon recruitment following agonist‐ induced phosphorylation of GPCRs by GRKs, the b‐arrestins, in addition to their role in the repression of signaling, may participate in the signal transduction acting as scaffolds for recruiting phosphatases and kinases, such as, for example, AKT and ERK (Hubbard and Hepler, 2006). In this perspective, b‐arrestins are essential for the AKT signaling involved in the Dopamine D2 receptor‐mediated behavior in mice (Beaulieu et al., 2004). Moreover, the action of b‐arrestins may not be limited to the role of a cytoplasmic adaptor in a signaling cascade, but may also intervene directly in gene regulation. b‐Arrestin1 has been found, after GPCR stimulation, in the nucleus as part of the nuclear complex formed, due to the promoter of target genes, by the transcription factor CREB and the histone acetyl transferase protein P300 (Kang et al., 2005).

2.6 Termination of NE Synaptic Transmission 2.6.1 Reuptake The synaptic activity of NE, as with the actions of all monoamines, is terminated by its active reuptake into the presynaptic neuron (known as uptake1) and/or glial cells. The uptake one mechanisms utilize Naþ/ Cl
‐dependent transporters (Lester et al., 1996). These transporters are members of a large family of Naþ/Cl
containing putative transmembrane domains, which control the concentration of the transmitter released into the intraplasmatic and extrasynaptic spaces via rapid reuptake into the nerve terminals, thus maintaining low concentration of the neurotransmitter at these sites (Masson et al., 1999). Since transporter velocity is increased by hyperpolarization of the membrane and is decreased by depolarization, autoreceptors or heteroreceptors affect the activity of the reuptake system by changing the membrane potential.

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The NE transporter (NET), like the serotonin or dopamine transporters, is a target for antidepressant drugs (ADs) and psychostimulant drugs. The NET is regulated by a wide range of intracellular signaling molecules such as PKC, c‐AMP/PKA, and CaM Kinase (Mandela and Ordway, 2006).

2.6.2 Metabolism The two enzymes that are important in the initial steps of metabolic transformation of NE and the other catecholamines are the monoamineoxidases (MAOs) and the catechol‐O‐methyltransferase (COMT). Two isoforms of MAO (MAO‐A and MAO‐B) are differentiated on the basis of substrate and inhibitor specificities. The preferential metabolizer of NE (and serotonin) is the MAO‐A whose specific inhibitor is clorgyline. Neurons contain both isoforms of MAOs, localized primarily in the outer membrane of mitochondria (Shih et al., 1999). Inhibitors of MAO cause an increase in the amount of monoamines stored and released from the nerve terminals, thus increasing their availability at the synapsis. COMT is bound to membranes and appears to be located principally in postsynaptic neurons. Both MAO and COMT enzymes, starting with either one of them, sequentially coordinate the degradation of NE. They generate aldehyde intermediates, which are reduced to 3,4‐dihydroxyphenyl‐glycol (DHPG) and 3‐methoxy‐4‐hydroxyphenylglycol (MHPG) by cytosolic aldehyde reductase or oxidized to vanyl‐mandelic acid (VMA) by mitochondrial aldehyde dehydrogenase (Eisenhofer et al., 2004).

3

Functional Noradrenergic System Neurotransmission

The postsynaptic effects of NE, at the cellular or neural circuit level, are essentially modulatory in nature. NE, rather than inducing simple inhibition or excitation, facilitates responses evoked in target cells by altering the ‘‘signal to noise ratio’’ (Servan‐Schreiber et al., 1990) of responses evoked by other afferents, both excitatory and inhibitory, thus enhancing synaptic transmission in target circuits (Woodward et al., 1991).

3.1 Brain Noradrenergic Neurotransmission and Arousal The involvement of the LC in the brain arousal system is essential for mediating attention in processes related to successful behavior. The LC is implicated in either selective (i.e. focused for rapid adaptative responses) or labile attention (Aston‐Jones et al., 1999). The LC innervates many more cerebral areas than any other brain nucleus, but it is also extremely specific in this innervation, which is most dense in areas associated with attention such as the parietal cortex, the pulvinar nucleus, and the superior colliculus. An overly focused or labile attention is a pathological behavior, which is a component of several neuropsychiatric diseases such as schizophrenia, ADHD, OCD, and depression.

3.1.1 Tonic Versus Phasic Excitation of NE Neurons NE is able to increase the evoked either excitatory or inhibitory activity while decreasing the spontaneous activity of its target neurons. NE plays a more specific role in regulating motivation and arousal, nonspecific aspects of behavior that are related in task‐related cognitive processes. In this aspect LC neurons display a stimulus‐specific activity similar to that of brainstem dopaminergic neurons responding selectively to stimuli that predict reward (Schultz et al., 1997). As observed in monkeys performing a visual discrimination task, LC neurons show short‐latency stimulus‐evoked (phasic) responses to target stimuli correlated with the corresponding behavioral responses but not to distractor stimuli or other task events. Thus, optimal behavioral performance is correlated with phasic activation of LC neurons. In contrast, periods of poor performance are associated with significantly higher tonic LC activity (Aston‐Jones et al., 1994). However, the LC tonic activity is necessary for an adaptative phasic activation. The LC tonic activity is high in waking, slow in sleeping, and almost absent during paradoxical sleep. The relationship between tonic and phasic activity related to behavioral performances corresponds graphically to an inverted‐U. The increase in

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phasic activation of the LC coupled with an intermediate tonic activity results in an excellent behavioral performance, whereas an increase or a decrease of the LC tonic activity diminishes the phasic activity with a concomitant reduction of the attentional performances. Thus, phasic LC discharge is related to behavioral responses and the level of LC tonic activity to behavioral performance (Aston‐Jones and Cohen, 2005).

3.2 Brain Noradrenergic Neurotransmission, Stress, and Depression Chronic stress has been implicated as a predisposing and/or triggering element for neuropsychiatric diseases such as major depression, bipolar disorder, and schizophrenia (Schildkraut, 1965) and has an important impact on the catecholaminergic system (Sabban and Kvetnansky, 2001) and in particular the NE‐ergic system (Morilak et al., 2005). Stress can be defined as any menace either real or perceived to the homeostasis and wellbeing of an organism. In the first case, stress is systemic or physiologic and the physical characteristics of the stressor require an immediate restorative response from the organism. Awareness or perception of the stimulus is not required to initiate the stress response. The second form is psychogenic stress, which depends upon perception, cognitive processing, and interpretation of the stimulus to confer upon it a stressful quality. In this case, the relative severity of the stressor and its physiologic impact is variable between individuals. The severity of stress presented by a stimulus, whether physiologic or psychogenic, has typically been defined in terms of the magnitude of the physiological response it elicits, for instance by measuring activation of the hormonal HPA stress axis, or of the peripheral sympathoadrenal autonomic response system. Stress in the experimental animal (restrain or foot shock) increases the firing in the LC and, in parallel, also the MHPG, which is a reliable index of the NE turnover. The inescapable stress not only induces a condition of ‘‘learned helplessness,’’ which is a preclinical correlate of depression, but also anxiety. Disregulation of NE‐ergic neurotransmission has been implicated in stress‐related psychiatric diseases such as depression, posttraumatic stress disorder (PTSD), and other anxiety disorders (Southwick et al., 1993; Sullivan et al., 1999; Leonard et al., 2001). Stress is able to deplete NE, and depletion of NE or repeated stress in rats upregulates TH in the LC (Melia and Duman, 1991; Melia et al., 1992). Elevated levels of TH have been found in the LC of victims of suicide (Ordway, 1997) and in patients with major depression as compared with normal controls (Zhu et al., 1999), whereas the NE transporter is downregulated in major depression (Klimek et al., 1997). Furthermore, the impairment of the NE‐ergic input from locus coeruleus to the forebrain limbic system may contribute to the symptoms of schizophrenia (Yamamoto and Hornykiewicz, 2004). In fact, these NE‐ergic projections are normally essential for screening and filtering the incoming sensory stimuli to discard irrelevant information (Archer, 1982). Alterations in NE‐ergic neurotransmission are important in the actions of many classes of antidepressant drugs (ADs), such as MAO inhibitors and tricyclic antidepressants, these latter blocking the reuptake of both NE and serotonin (Nelson, 1999). Serotonin in particular has been heralded as the main player in depression, since inhibition of its synthesis counteracts the effects of antidepressants, and selective serotonin reuptake inhibitors (SSRIs) are highly effective in the regulation of mood (Fuller et al., 1995). However, the observation that the extremely selective NE reuptake inhibitor reboxetine is an effective and powerful antidepressant has reactualized the role of NE in depression (Schatzberg, 2000). Activation of NE release in the limbic forebrain by acute stressors may facilitate anxiety‐like behavioral responses, making anxiety a prominent component of depression. The new generation of dual uptake inhibitors, as well as selective NE reuptake inhibitors, alleviate depressed mood, social withdrawal, and other symptoms of depression including anxiety (Nelson, 1999; Versiani et al., 2002; Ferguson et al., 2003; Morilak et al., 2005).

4

Genetics of Noradrenergic Neurotransmission

The role played by central NE‐ergic neurotransmission in behavior and its implication in related disturbances have prompted, with the advent of positional cloning, the investigation of linkage or association to neuropsychiatric diseases of the genes encoding the components of this system. These studies have been hampered by the genetic complexity inherent to psychiatric diseases, which are characterized by

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heterogeneity, polygenicity, and the interplay between genetic and environmental factors in such a way that no clear‐cut mutations but only predisposing polymorphism are implicated in their etiology. Taking into account these caveats, the most relevant results in this domain have been obtained for the TH, DBH, MAO, and COMT genes.

4.1 Tyrosine Hydroxylase The TH gene, located in the chromosome 11p15, encodes the rate‐limiting enzyme in the synthesis of catecholamines, and is thus a strong candidate gene for neuropsychiatric diseases (Mallet, 1996). A first report showing a genetic linkage in the Amish population between bipolar disorder and markers at the chromosomal region that contains the TH locus also put forward the TH as a ‘‘positional’’ candidate gene (Egeland et al., 1987). This result was questioned because the lod score method utilized did not take into account genetic heterogeneity that characterizes bipolar disorder and other complex diseases (Hodgkinson et al., 1987), and neither further genetic analysis of the Amish nor studies of other populations confirmed this initial result (Detera‐Wadleigh et al., 1987; Kelsoe et al., 1989; Ginns et al., 1992; Gerhard et al., 1994; Gershon et al., 1996; Ginns et al., 1996). However, other studies finding significant linkage between markers at the TH locus and bipolar disorder maintained the implication of the TH gene in this disease and strengthened the case for genetic heterogeneity (Pakstis et al., 1991; Byerley et al., 1992; Lim et al., 1993; Gurling et al., 1995; Smyth et al., 1996; Malafosse et al., 1997). In the first association study of the TH gene, a significant genetic association was found between restriction fragment polymorphism markers at the TH locus and bipolar disorder in a French population sample (Leboyer et al., 1990), a result that was not always replicated in other studies (Korner et al., 1990, 1994; Gill et al., 1991; Inayama et al., 1993; Kawada et al., 1995). Another association analysis was conducted using the more informative microsatellite HUMTH01 marker in order to further investigating the implication of the TH gene in the genetic predisposition to bipolar disorder. This microsatellite is a polymorphic polypyrimidine sequence localized in the first intron of the TH gene and is characterized by a core (TCAT)n tetranucleotide repeat iterated usually between 5 and 10 times (Polymeropoulos et al., 1991; Puers et al., 1993; Brinkmann et al., 1996). In a new sample of French case–controls, a significant genotypic association was found between the HUMTH01 and bipolar disorder as well as familial history of bipolar disorder and/or delusive symptoms during manic or depressive episodes (Meloni et al., 1995a). Moreover, a rare allele of the HUMTH01 microsatellite was significantly associated with schizophrenia in two different ethnic samples from Normandy in northwestern France and the Sousse region of eastern Tunisia (Meloni et al., 1995b). Several further studies inspired by these results have been inconclusive for association (Cavazzoni et al., 1996; Souery et al., 1996, 1999; Turecki et al., 1997; Burgert et al., 1998; Jonsson et al., 1998) or have replicated the positive association between this microsatellite and both bipolar disorder (Perez de Castro et al., 1995; Lobos and Todd, 1997; Serretti et al., 1998a, b; Furlong et al., 1999; Chiba et al., 2000) and schizophrenia (Wei et al., 1995, 1997; Kurumaji et al., 2001). It is noteworthy that the HUMTH01 microsatellite is associated with other physiological or behavioral traits such as personality (Persson et al., 2000), longevity (De Benedictis et al., 1998), symptoms of alcohol withdrawal (Sander et al., 1998), hypertension (Jindra et al., 2000), and stress response parameters in twins (Zhang et al., 2004), but also directly with catecholamine neurotransmission. This is shown by measuring catecholamine metabolite levels in cerebrospinal fluid (Jonsson et al., 1996) or in plasma (Wei et al., 1995, 1997) as an indirect index of their turnover in the brain. Moreover, in a clinical study in the original Normandy sample the schizophrenic patients bearing the rare allele associated with schizophrenia presented significantly lower plasma concentrations of the catecholaminergic metabolites MHPG and HVA, which are indices of central NE‐ergic and DA‐ergic function, respectively, as compared with patients bearing other alleles (Thibaut et al., 1997). These results suggest a functional link between allelic variations at the HUMTH01 marker and TH activity. Indeed, the (TCAT)n motif of this microsatellite differs by only one nucleotide from the consensus AP1 sequence (TGATTCA) present in the rat and the human TH gene (Icard Liepkalns et al., 1992), a sequence that is specifically recognized by transcription factors of the Fos and Jun proto‐onco‐gene families

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(Sassone‐Corsi et al., 1988). Moreover, a less polymorphic HUMTH01 repeated sequence is conserved at its orthologous position in the first intron of the TH gene in several non‐human primate species (Meyer et al., 1995), suggesting that this motif may be an evolutionary conserved regulatory element that has expanded in the human lineage. Therefore, the functional role of the HUMTH01 microsatellite was assessed to investigate the biological significance of the genetic association findings. Indeed, the alleles of this microsatellite enhanced transcription when placed upstream from a minimal promoter driving the expression of a luciferase‐reporter gene. Moreover, these repeated sequences interacted specifically with factors of the fos/jun type and with an even higher affinity with other nuclear proteins (Meloni et al., 1998). Subsequently, ZNF191, a zinc finger protein, and HBP1, a HMG box transcription factor, were identified as the proteins specifically binding the TCAT motif. Interestingly, the specific binding of ZNF191 to the HUMTH01 sequence was correlated in a quantitative fashion to the number of TCAT repeats. Moreover, in vitro experiments with a TH‐reporter gene construct established that the HUMTH01 microsatellite regulates the TH gene expression by a quantitative silencing effect that correlates with the number of repetitions of the (TCAT) motif (Albanese et al., 2001). Thus, the HUMTH01 sequence may participate in the transcriptional regulation of the TH gene by modulating its expression in a quantitative fashion. It is noteworthy that HBP1 was characterized as a chromatin remodeling factor, which binds specifically a TCAT short repeat in the locus control region of the CD2 gene (Zhuma et al., 1999). Moreover, a polypyrimidine trait similar to the TCAT repeated motif has been shown to regulate the expression of the CD30 gene (Croager et al., 2000). Finally, a TCAT stretch is the only difference that characterizes the regulatory region of the vasopressin receptor gene that accounts for a different distribution of the gene product in a brain region of two different species of voles. This single difference is responsible for completely different mating and parental behavior between species (Young et al., 1999). Since the (TCAT)n polymorphic sequence is widespread in the genome and present in several genes, it may provide a molecular basis for the modulation of gene expression relevant to the genetics of quantitative traits.

4.2 Dopamine b Hydroxylase The DBH gene is located on chromosome 9q34 (Craig et al., 1988) and encodes the enzyme that catalyzes the conversion of DA to NE. Mutations in the DBH gene result in a lack of sympathetic NE‐ergic function and orthostatic hypotension (Garland et al., 2002; Deinum et al., 2004). The DBH enzyme is localized within the soluble and membrane fractions of secretory catecholamine‐containing vesicles of NE‐ergic and adrenergic cells. These two forms originate from optional cleavage of the signal peptide. The form retaining the signal peptide is completely associated with the membrane, whereas the cleaved form is mostly soluble (Houhou et al., 1995). The soluble form of the enzyme is secreted into the circulation from nerve terminals allowing for assaying its activity in plasma or serum. DBH presents stable differences in enzymatic activity that appear to be genetically determined (Stolk et al., 1982). Several polymorphisms in the human gene have been implicated in these variations. A G/T polymorphism at the nucleotide 910 of the coding sequence results in a change of amino acid residue 304 between Ala (A) and Ser (S) (DBH/A and DBH/S). The resulting proteins have similar kinetic constants, but DBH/S has a homospecific activity that is about 1/13 lower than that of DBH/A (Ishii et al., 1991). In European, African, and several other populations the DBH/ A allele is the most common with allele frequencies greater than 0.80 in each sample and significant heterogeneity in allele frequency across population groups (Cubells et al., 1997). These allelic differences cannot alone account for the differences in the activity of DBH in blood since circulating DBH concentrations also vary considerably in the general population. Recently, a novel polymorphism (
1021 C/T) in the 50 promoter region of the DBH gene was shown to strongly influence plasma DBH activity, accounting for 35–52% of its variation in different populations (Zabetian et al., 2001; Kohnke et al., 2002). A further study showed that ten biallelic markers in a 10 Kb surrounding the
1021C/T polymorphisms were all associated with plasma DBH activity and that this association was strongly correlated with the degree of Linkage Disequilibrium between each marker and the
1021C/T polymorphism (Zabetian et al., 2003). Another association was found between a DBH TaqI polymorphism and plasma metabolites of catecholamines

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(Wei et al., 1998). Other polymorphic variants in the DBH gene are represented by a GT dinucleotide microsatellite, a single‐base, 444 g/a, substitution at the 30 end of DBH exon 2 and a diallelic variant, DBH50 ‐ins/del, located approximately 3 kb 50 to the DBH transcriptional start site. All these markers, which are in linkage disequilibrium, were also associated with plasma DBH activity (Wei et al., 1997; Cubells et al., 1998, 2000; Jo¨nsson et al., 2004). Moreover, the 444 g/a marker was also associated with differences in DBH concentration in the CSF (Cubells et al., 1998; Zabetian et al., 2003). Psychiatric genetic studies using the polymorphisms at the DBH gene have shown a significant association between the DBH TaqI polymorphism and ADHD (Daly et al., 1999). Also, albeit it did not reach statistical significance, the DBH GT repeat four allele, which is associated with high serum levels of DBH, occurred more frequently in the ADHD group than controls (Mu¨ller Smith et al., 2003). However, other groups failed in replicating these results either with the same (Wigg et al., 2002) or other markers (Hawi et al., 2003). A positive association was shown between the DBH 50 ‐444a haplotype and cocaine‐ induced paranoia (Cubells et al., 2000) as well as a lack of response to antipsychotic drug treatment in schizophrenic patients (Yamamoto et al., 2003). These results may indicate that the DBH gene is indirectly involved in schizophrenia as a modulatory factor of psychotic symptoms, severity of the disorder, and therapeutic response to neuroleptic drugs.

4.3 Mono‐Amine‐Oxydase The MAO‐A and MAO‐B genes are situated on the X chromosome at Xp11.23–11.4 and result from the duplication of a common ancestral gene. In humans both genes are deleted in patients with Norrie’s disease, a rare X‐linked recessive neurological disorder characterized by blindness, hearing loss, and mental retardation (Lan et al., 1989). A point deletion resulting in a complete MAO‐A inactivation was linked to abnormally aggressive behavior in the males of a Dutch family (Brunner et al., 1993). Conversely, MAO‐A knock‐out mice show an increased aggressivity in males, which is related to increased levels of NE and serotonin during development and result in brain structural changes (Cases et al., 1995). A functional polymorphism located in the MAO‐A gene promoter 1.2kb upstream of the encoding sequence consists of a 30bp repeated sequence present in 3, 3.5, 4, or 5 copies. This polymorphism displays significant variations in allele frequencies across ethnic groups and is able to affect the transcriptional activity of the MAO‐A gene promoter (Sabol et al., 1998). Genetic studies with this polymorphism have found that the high‐activity MAO‐A gene promoter alleles were associated with panic disorder (Deckert et al., 1999) and major depressive disorder (Schulze et al., 2000) in females, whereas the low‐activity alleles were associated with schizophrenia in males (Jonsson et al., 2003). Other studies have yielded negative results for panic disorders (Hamilton et al., 2000), schizophrenia (Syagailo et al., 2001; Fan et al., 2004), and mood disorders (Kirov et al., 1999; Kunugi et al., 1999; Jorm et al., 2000; Syagailo et al., 2001; Huang et al., 2004). However, more probant results have been found when genetic studies have taken into account an environmental component. A leading study has shown that this functional polymorphism can modulate the association between childhood maltreatment and subsequent antisocial behavior. In males, who have only one copy of the X chromosome, the MAO‐A functional polymorphism confers either a high or a low‐activity genotype. The low‐activity MAO‐A genotype is associated with antisocial behavior in up to 85% of a cohort of males who had been severely maltreated in their childhood but not in boys who had suffered little or no abuse. In contrast, the high‐activity MAO‐A genotype has a protective effect from developing antisocial behavior in maltreated children. In women a similar trend for association between MAO‐A genotype, antisocial behavior and child maltreatment, was present, but the interpretation of this result was hindered by the presence of two X chromosomes with sometimes heterozygous low/high MAO‐A activity alleles, one of which is randomly inactivated. However, taken together, these results show a clear influence of the MAO‐A genotype in the behavioral effects of an environmental factor and may help in understanding the marked differences in the frequency of antisocial behavior between sexes (Caspi et al., 2002). The association between the lower expression MAO‐A genotype and antisocial behavior consequent to childhood maltreatment has been replicated

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(Foley et al., 2004), and this risk genotype has also been associated with impulsive traits in males who have experienced early abuse (Huang et al., 2004) and with pathological gambling always in males (Ibanez et al., 2000).

4.4 Catechol‐O‐Methyltransferase The COMT gene, localized to chromosome 22q11.1–q11.2, encodes a soluble (S‐COMT) and a membrane‐ bound (MB‐COMT) form of the enzyme, the latter characterized by an additional 50 amino acids at the N‐terminal (Bertocci et al., 1991; Grossman et al., 1992). The two length variants of the COMT are expressed from two mRNA transcripts: a long mRNA, which is able to translate both S‐COMT and MB‐COMT from two different initiation sites, and a short mRNA producing S‐COMT only. The long mRNA and the larger MB‐COMT are predominant in the brain whereas the short mRNA and the S‐COMT prevail in the other tissues (Tenhunen et al., 1994; Lundstrom et al., 1995). The COMT enzymatic activity shows high, intermediate, and low rates consistent with inheritance of two codominant alleles (Weinshilboum, 1978). This difference in enzyme activity is independent of protein length variations but is caused by an amino acid substitution. A G/A polymorphism in exon 4 at position 472 in the long mRNA, and 322 in the short mRNA, results in a Val to Met amino acid change at codon 158 of MB‐COMT and codon 108 of S‐COMT. The G (Val) allele encodes the thermostable, high‐activity form of the enzyme, whereas the A (Met) allele encodes the thermolabile, low‐activity variant that exhibits a 3‐ to 4‐fold decrease in the enzymatic activity level (Lotta et al., 1995; Lachman et al., 1996a, b). The G (Val) and A (Met) alleles correspond also to the absence or presence, respectively, of a NlaIII polymorphic restriction site that allows for easily genotyping the functional variations (Karayiorgou et al., 1998). COMT is an obvious a priori candidate gene for neuropsychiatric disorders that involve dopaminergic or NE‐ergic systems (Palmatier et al., 1999) but also a strong positional candidate gene for schizophrenia because of its chromosomal location in the locus of the velocardiofacial (VCF) syndrome. Microdeletions of 22q11 are associated with VCF syndrome which is characterized by congenital abnormalities, learning difficulties, and psychosis in up to one third of patients. Conversely, the deletion is also 80‐fold more common in patients with psychosis than the normal population (Sugama et al., 1999). Both linkage and association studies have implied that chromosome 22q11 is a locus for schizophrenia (Pulver et al., 1994a, b; Karayiorgou et al., 1995; Karayiorgou and Gogos, 1997). The case–control association approach has consequently been used to study the role of COMT in schizophrenia and other psychiatric diseases, mostly using the Val108/158Met polymorphism. Positive associations have been found between COMT and schizophrenia (Ohmori et al., 1998; de Chaldee et al., 1999), violence in schizophrenia (Lachman et al., 1998), bipolar disorder (Li et al., 1997; Mynett‐Johnson et al., 1998), unipolar disorder (Ohara et al., 1998), bipolar disorder or ADHD in VCF syndrome patients (Lachman et al., 1996a), OCD (Karayiorgou et al., 1997), drug abuse (Vandenbergh et al., 1997), and Parkinson’s disease (Kunugi et al., 1997a). However, other studies have excluded a major contribution of the COMT gene to schizophrenia (Daniels et al., 1996; Chen et al., 1997; Strous et al., 1997; Karayiorgou et al., 1998; Wei and Hemmings, 1999), bipolar disorder (Craddock et al., 1997; Gutierrez et al., 1997; Kunugi et al., 1997b; Lachman et al., 1997; Geller and Cook, 2000), ADHD or bipolar disorder in VCF syndrome patients (Lachman et al., 1996b), substance abuse and violence (Vandenbergh et al., 1997; Lachman et al., 1998), as well as Parkinson’s disease (Hoda et al., 1996; Syvanen et al., 1997; Xie et al., 1997). These conflicting results have prompted a meta‐analysis indicating that the COMT Met allele that characterizes the instable form of the enzyme with low‐activity phenotype is not associated with schizophrenia (Lohmueller et al., 2003). However, a new association study conducted in a genetically homogeneous population yielded a highly significant association between a COMT haplotype and schizophrenia (Shifman et al., 2003). This study is the largest case/control analysis in schizophrenia that has been reported with more than 700 patients and 4,000 control subjects. Genotyping was conducted using 12 SNPs, comprising the Val108/158Met polymorphism, across the COMT gene, and haplotypes with seven of these SNPs were established in the large sample of an Israeli Ashkenazi Jewish population. This population has the advantage of presenting a founder effect that allows for reducing genetic heterogeneity thus increasing gene effect and avoiding false‐positive results due to population stratification.

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The association between schizophrenia and the Val108/158Met polymorphism was moderate, but extremely high levels of statistical significance were attained when this marker was analyzed as part of a haplotype including two other noncoding SNPs that were more significantly associated with schizophrenia. Moreover, one of these polymorphisms represented a higher risk factor essentially for women than men, hinting at a possible sex‐specific genetic component in schizophrenia. These results confirmed a complex association of the COMT locus to schizophrenia and suggested that other functional variants besides the Val108/158Met polymorphism are likely to be involved in susceptibility to the disease (Shifman et al., 2003). A significant association was also found between bipolar disorder and both the allele and haplotype of the COMT gene and they were previously found to be associated with schizophrenia (Shifman et al., 2004). In addition to these association studies, the role of COMT in schizophrenia and other neuropsychiatric diseases is further supported by functional genetic studies that have essentially focused on the Val108/ 158Met polymorphism. A large amount of experimental data suggests that heritable abnormalities of prefrontal cortex function are a prominent feature of schizophrenia (Grace, 1991, 1993; Moore et al., 1999). COMT may constitute a major contributor to these abnormalities by virtue of its unique role in regulating catecholamine‐mediated prefrontal information processing, since COMT inhibitors can improve working memory in both rodents and humans (Weinberger et al., 2001). In this perspective, a study combining a genetic and a functional approach has shown that the Val allele of the Val108/158Met polymorphism that characterizes the high‐activity form of the COMT occurs at higher rates in both schizophrenics and their unaffected siblings. Moreover, patients and siblings bearing this allele performed poorly on the Wisconsin card sorting test (a neuropsychological test of frontal lobe function for working memory) and manifested inefficient brain activation as assessed by functional magnetic resonance imaging (fMRI) (Egan et al., 2001). Interestingly, amphetamine, a drug that increases catecholaminergic neurotransmission, enhances the efficiency of prefrontal cortex function as assayed with fMRI during a working memory task in subjects with the high‐activity val/val genotype but not in subjects with the low‐activity met/met genotype (Mattay et al., 2003). Moreover, this polymorphism is also associated with personality traits, as assessed by the tridimensional personality questionnaire (Benjamin et al., 2000). In addition, homozygosity for the Met allele is associated, particularly in schizophrenic patients, with lower frontal P300 amplitudes, which is an index of catecholaminergic efficacy in reducing noise during information processing (Gallinat et al., 2003). In agreement with these findings and with the results of the association studies in Ashkenazi Jews (Shifman et al., 2003, 2004), the analysis of the allele‐specific expression using mRNA from human brains indicated that the haplotype implicated in schizophrenia and bipolar disorder is associated with lower expression of COMT mRNA (Bray et al., 2003). These findings suggest that the COMT Val allele impairs prefrontal cognition and physiology and, by virtue of this effect, may condition some pathological features of schizophrenia thus contributing, with other sequence variations at the COMT locus, to the increase of the risk for schizophrenia or other psychiatric disorder characterized by impaired frontal cortex functioning.

5

Conclusions

The brain NE‐ergic system plays a pivotal role in integrating and fine‐tuning the adaptative responses to basic arousal and stress‐generated stimuli. Anatomical, functional, and genetic studies may further contribute to the understanding of how the mechanism underlying these responses may intervene in normal and pathological behavior and yield new entries for therapeutic interventions in related psychiatric diseases.

References Albanese V, Biguet NF, Kiefer H, Bayard E, Mallet, J, et al. 2001. Quantitative effects on gene silencing by allelic variation at a tetranucleotide microsatellite. Hum Mol Genet 10: 1785-1792. Archer T. 1982. The role of noradrenaline in learned behaviors: Studies using DSP4. Scand J Psychol Suppl 1: 61-71.

Aston‐Jones G, Cohen, JD. 2005. An integrative theory of locus coeruleus‐norepinephrine function: Adaptive gain and optimal performance. Annu Rev Neurosci 28: 403-450. Aston‐Jones G, Rajkowski J, Cohen J. 1999. Role of locus coeruleus in attention and behavioral flexibility. Biol Psychiatry 46: 1309-1320.

Molecular genetics of brain noradrenergic neurotransmission Aston‐Jones G, Rajkowski J, Kubiak P, Alexinsky T. 1994. Locus coeruleus neurons in monkey are selectively activated by attended cues in a vigilance task. J Neurosci 14: 4467-4480. Beaulieu J‐M, Sotnikova TD, Yao W.‐D, Kockeritz L, Woodgett JR, et al. 2004. Lithium antagonizes dopamine‐ dependent behaviors mediated by an AKT/glycogen synthase kinase 3 signaling cascade. Proc Natl Acad Sci USA 101: 5099-5104. Benjamin J, Osher Y, Kotler M, Gritsenko I, Nemanov L, et al. 2000. Association between tridimensional personality questionnaire (TPQ) traits and three functional polymorphisms: Dopamine receptor D4 (DRD4), serotonin transporter promoter region (5‐HTTLPR) and catechol‐ O‐methyltransferase (COMT). Mol Psychiatry 5: 96-100. Berridge M. 1993. Cell signalling. A tale of two messengers. Nature 365: 388-389. Berridge CW, Waterhouse BD. 2003. The locus coeruleus‐ noradrenergic system: Modulation of behavioral state and state‐dependent cognitive processes. Brain Res Rev 42: 33-84. Bertocci B, Miggiano V, Da Prada M, Dembic Z, Lahm, HW, et al. 1991. Human catechol‐O‐methyltransferase: Cloning and expression of the membrane‐associated form. Proc Natl Acad Sci USA 88: 1416-1420. Boehm S, Kubista, H. 2002. Fine tuning of sympathetic transmitter release via ionotropic and metabotropic presynaptic receptors. Pharmacol Rev 54: 43-99. Bray N, Buckland P, Williams N, Williams H, Norton N, et al. 2003. A haplotype implicated in schizophrenia susceptibility is associated with reduced COMT expression in human brain. Am J Hum Genet 73: 152-161. Brinkmann B, Sajantila A, Goedde H, Matsumoto H, Nishi K, et al. 1996. Population genetic comparisons among eight populations using allele frequency and sequence data from three microsatellite loci. Eur J Hum Genet 4: 175-182. Brunet J‐F, Pattyn A. 2002. Phox2 genes—from patterning to connectivity. Curr Opin Genet Dev 12: 435-440. Brunner HG, Nelen M, Breakfield XO, Ropers HH, Van Oost B. 1993. Abnormal behavior associated with a point mutation in the structural gene for monoamine oxidase A. Science 262: 578-580. Burgert E, Crocq MA, Bausch E, Macher JP, Morris‐ Rosendahl DJ. 1998. No association between the tyrosine hydroxylase microsatellite marker HUMTH01 and schizophrenia or bipolar I disorder. Psychiatr Genet 8: 45-48. Byerley W, Plaetke R, Hoff M, Jensen S, Holik J, et al. 1992. Tyrosine hydroxylase gene not linked to manic‐depression in seven of eight pedigrees. Hum Hered 42: 259-263. Bylund D. 1992. Subtypes of alpha 1‐ and alpha 2‐adrenergic receptors. FASEB J 6: 832-839. Bylund, DB. 2005. Alpha‐2 adrenoceptor subtypes: Are more better? 144: 159‐160.

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Bylund D, Eikenberg D, Hieble J, Langer S, Lefkowitz R, et al. 1994. International union of pharmacology nomenclature of adrenoceptors. Pharmacol Rev 46: 121-136. Cases O, Self I, Grimsby J, Gaspar P, Chen K, et al. 1995. Aggressive behavior and altered amounts of brain serotonin and norepinephrine in mice lacking MAOA. Science 268: 1763-1766. Caspi A, McClay J, Moffitt TE, Mill J, Martin J, et al. 2002. Role of genotype in the cycle of violence in maltreated children. Science 297: 851-854. Cavazzoni P, Alda M, Turecki G, Rouleau G, Grof E, et al. 1996. Lithium‐responsive affective disorders: No association with the tyrosine hydroxylase gene. Psychiatry Res 64: 91-96. de Chaldee M, Laurent C, Thibaut F, Martinez M, Samolyk D, et al. 1999. Linkage disequilibrium on the COMT gene in French schizophrenics and controls. Am J Med Genet 88: 452-457. Chen CH, Lee YR, Wei FC, Koong FJ, Hwu HG, et al. 1997. Association study of NlaIII and MspI genetic polymorphisms of catechol‐O‐methyltransferase gene and susceptibility to schizophrenia. Biol Psychiatry 41: 985-987. Chiba M, Suzuki S, Hinokio Y, Hirai M, Satoh Y, et al. 2000. Tyrosine hydroxylase gene microsatellite polymorphism associated with insulin resistance in depressive disorder. Metabolism 49: 1145-1149. Craddock N, Spurlock G, McGuffin P, Owen MJ, Nosten‐ Bertrand M, et al. 1997. No association between bipolar disorder and alleles at a functional polymorphism in the COMT gene. Br J Psychiatry 170: 526-528. Craig SP, Buckle VJ, Lamouroux A, Mallet J, Craig IW. 1988. Localization of the human dopamine beta hydroxylase (DBH) gene to chromosome 9q34. Cytogenet Cell Genet 48: 48-50. Croager EJ, Gout AM, Abraham LJ. 2000. Involvement of Sp1 and microsatellite repressor sequences in the transcriptional control of the human CD30 gene. Am J Pathol 156: 1723-1731. Cubells JF, van Kammen DP, Kelley ME, Anderson GM, O’Connor DT, et al. 1998. Dopamine beta‐hydroxylase: Two polymorphisms in linkage disequilibrium at the structural gene DBH associate with biochemical phenotypic variation. Hum Genet 102: 533-540. Cubells JF, Kobayashi K, Nagatsu T, Kidd KK, Kidd JR, et al. 1997. Population genetics of a functional variant of the dopamine beta‐hydroxylase gene (DBH). Am J Med Genet 74: 374-379. Cubells JF, Kranzler HR, McCance‐Katz E, Anderson GM, Malison RT, et al. 2000. A haplotype at the DBH locus, associated with low plasma dopamine beta‐hydroxylase activity, also associates with cocaine‐induced paranoia. Mol Psychiatry 5: 56-63. Daly G, Hawi Z, Fitzgerald M, Gill M. 1999. Mapping susceptibility loci in attention deficit hyperactivity disorder:

141

142

6

Molecular genetics of brain noradrenergic neurotransmission

Preferential transmission of parental alleles at DAT1, DBH and DRD5 to affected children. Mol Psychiatry 4: 192-196. Daniels JK, Williams NM, Williams J, Jones LA, Cardno AG, et al. 1996. No evidence for allelic association between schizophrenia and a polymorphism determining high or low catechol‐O‐methyltransferase activity. Am J Psychiatry 153: 268-270. De Benedictis G, Carotenuto L, Carrieri G, De Luca M, Falcone E, et al. 1998. Gene/longevity association studies at four autosomal loci (REN, THO, PARP, SOD2). Eur J Hum Genet 6: 534-541. Deckert J, Catalano M, Syagailo Y, Bosi M, Okladnova O, et al. 1999. Excess of high activity monoamine oxidase A gene promoter alleles in female patients with panic disorder. Hum Mol Genet 8: 621-624. Deinum J, Steenbergen‐Spanjers GCH, Jansen M, Boomsma F, Lenders JWM, et al. 2004. DBH gene variants that cause low plasma dopamine {beta} hydroxylase with or without a severe orthostatic syndrome. J Med Genet 41: e38. Detera‐Wadleigh S, Berrettini W, Goldin L, Boorman D, Anderson S, et al. 1987. Close linkage of c‐Harvey‐ras‐1 and the insulin gene to affective disorder is ruled out in three North American pedigrees. Nature 325: 806-808. Egan MF, Goldberg TE, Kolachana BS, Callicott JH, Mazzanti CM, et al. 2001. Effect of COMT val108/158 met genotype on frontal lobe function and risk for schizophrenia. Proc Natl Acad Sci USA 98: 6917-6922. Egeland JA, Gerhard DS, Pauls DL, Sussex JN, Kidd KK, et al. 1987. Bipolar affective disorders linked to DNA markers on chromosome 11. Nature 325: 783-787. Eisenhofer G, Kopin IJ, Goldstein DS. 2004. Catecholamine metabolism: A contemporary view with implications for physiology and medicine. Pharmacol Rev 56: 331-349. Fan J‐B, Yang M‐S, Tang J‐X, He L, Xing YL, et al. 2004. Family‐based association study of the functional monoamine oxidase a gene promoter polymorphism and schizophrenia. Schizophr Res 67: 107-109. Ferguson SSG. 2001. Evolving concepts in G protein‐coupled receptor endocytosis: The role in receptor desensitization and signaling. Pharmacol Rev 53: 1-24. Ferguson J, Wesnes K, Schwartz G. 2003. Reboxetine versus paroxetine versus placebo: Effects on cognitive functioning in depressed patients. Int Clin Psychopharmacol 18: 9-14. Fields TA, Casey PJ. 1997. Signalling functions and biochemical properties of pertussis toxin‐resistant G‐proteins. Biochem J 321: 561-571. Foley DL, Eaves LJ, Wormley B, Silberg JL, Maes HH, et al. 2004. Childhood adversity, monoamine oxidase a genotype, and risk for conduct disorder. Arch Gen Psychiatry 61: 738-744. Fuller R, Hemrick‐Luecke S, Littlefield E, Audia J. 1995. Comparison of desmethylsertraline with sertraline as a

monoamine uptake inhibitor in vivo. Prog Neuropsychopharmacol Biol Psychiatry 19: 135-149. Furlong RA, Rubinsztein JS, Ho L, Walsh C, Coleman TA, et al. 1999. Analysis and metaanalysis of two polymorphisms within the tyrosine hydroxylase gene in bipolar and unipolar affective disorders. Am J Med Genet 88: 88-94. Gallinat J, Bajbouj M, Sander T, Schlattmann P, Xu K, et al. 2003. Association of the G1947A COMT (Val108/158Met) gene polymorphism with prefrontal P300 during information processing. Biol Psychiatry 54: 40-48. Garland EM, Hahn MK, Ketch TP, Keller NR, Kim CH, et al. 2002. Genetic Basis of clinical catecholamine disorders. Ann NY Acad Sci 971: 506-514. Geller B, Cook Jr., EH. 2000. Ultradian rapid cycling in prepubertal and early adolescent bipolarity is not in transmission disequilibrium with val/met COMT alleles. Biol Psychiatry 47: 605-609. Gerhard D, LaBuda M, Bland S, Allen C, Egeland J, et al. 1994. Initial report of a genome search for the affective disorder predisposition gene in the old order amish pedigrees: Chromosomes 1 and 11. Am J Med Genet 54: 398-404. Gershon E, Goldin L, Badner J, Berrettini W. 1996. Detection of linkage to affective disorders in the catalogued amish pedigrees: A reply to Pauls et al. Am J Hum Genet 58: 1381-1385. Gether U. 2000. Uncovering molecular mechanisms involved in activation of G protein‐coupled receptors. Endocr Rev 21: 90-113. Gill M, Castle D, Hunt N, Clements A, Sham P, et al. 1991. Tyrosine hydroxylase polymorphisms and bipolar affective disorder. J Psychiatr Res 25: 179-184. Ginns E, Egeland J, Allen C, Pauls D, Falls K, et al. 1992. Update on the search for DNA markers linked to manic‐ depressive illness in the old order amish. J Psychiatr Res 26: 305-308. Ginns E, Ott J, Egeland J, Allen C, Fann C, et al. 1996. A genome‐wide search for chromosomal loci linked to bipolar affective disorder in the old order amish. Nat Genet 12: 431-435. Goridis C, Rohrer H. 2002. Specification of catecholaminergic and serotonergic neurons. Nat Rev Neurosci 3: 531-541. Grace AA. 1991. Phasic versus tonic dopamine release and the modulation of dopamine system responsivity: A hypothesis for the etiology of schizophrenia. Neuroscience 41: 1-24. Grace AA. 1993. Cortical regulation of subcortical dopamine systems and its possible relevance to schizophrenia. J Neural Transm Gen Sect 91: 111-134. Grossman MH, Emanuel BS, Budarf ML. 1992. Chromosomal mapping of the human catechol‐O‐methyltransferase gene to 22q11.1–q11.2. Genomics 12: 822-825.

Molecular genetics of brain noradrenergic neurotransmission Gurling H, Smyth C, Kalsi G, Moloney E, Rifkin L, et al. 1995. Linkage findings in bipolar disorder. Nat Genet 10: 8-9. Gutierrez B, Bertranpetit J, Guillamat R, Valles V, Arranz MJ, et al. 1997. Association analysis of the catechol‐ O‐methyltransferase gene and bipolar affective disorder. Am J Psychiatry 154: 113-115. HamiltonS, Slager S, Heiman G, Haghighi F, Klein D, et al. 2000. No genetic linkage or association between a functional promoter polymorphism in the monoamine oxidase‐ A gene and panic disorder. Mol Psychiatry 5: 465-466. Hawi Z, Lowe N, Kirley A, Gruenhage F, Nothen M, et al. 2003. Linkage disequilibrium mapping at DAT1, DRD5 and DBH narrows the search for ADHD susceptibility alleles at these loci. Mol Psychiatry 8: 299-308. Hoda F, Nicholl D, Bennett P, Arranz M, Aitchison KJ, et al. 1996. No association between Parkinson’s disease and low‐activity alleles of catechol‐O‐methyltransferase. Biochem Biophys Res Commun 228: 780-784. Hodgkinson S, Sherrington R, Gurling H, Marchbanks R, Reeders S, et al. 1987. Molecular genetic evidence for heterogeneity in manic depression. Nature 325: 805-806. Houhou L, Lamouroux A, Faucon Biguet N, Mallet J. 1995. Expression of human dopamine b‐hydroxylase in mammalian cells infected by recombinant vaccinia virus. J Biol Chem 270: 12601-12606. Huang Y, Cate S, Battistuzzi C, Oquendo M, Brent D, et al. 2004. An association between a functional polymorphism in the monoamine oxidase a gene promoter, impulsive traits and early abuse experiences. Neuropsychopharmacology 29: 1498-1505. Hubbard KB, Hepler JR. 2006. Cell signalling diversity of the Gq[alpha] family of heterotrimeric G proteins. Cellular Signalling 18: 135-150. Ibanez A, Perez de Castro I, Fernandez‐Piqueras J, Blanco C, Saiz‐Ruiz J. 2000. Pathological gambling and DNA polymorphic markers at MAO‐A and MAO‐B genes. Mol Psychiatry 5: 105-109. Icard Liepkalns C, Faucon Biguet N, Vyas S, Robert JJ, Sassone Corsi P, et al. 1992. AP‐1 complex and c‐fos transcription are involved in TPA provoked and trans‐synaptic inductions of the tyrosine hydroxylase gene: Insights into long‐term regulatory mechanisms. J Neurosci Res 32: 290-298. Inayama Y, Yoneda H, Sakai T, Ishida T, Kobayashi S, et al. 1993. Lack of association between bipolar affective disorder and tyrosine hydroxylase DNA marker. Am J Med Genet 48: 87-89. Ishii A, Kobayashi K, Kiuchi K, Nagatsu T. 1991. Expression of two forms of human dopamine‐beta‐hydroxylase in COS cells. Neurosci Lett 125: 25-28. Jindra A, Jachymova M, Horky K, Peleska J, Umnerova V, et al. 2000. Association analysis of two tyrosine hydroxylase

6

gene polymorphisms in normotensive offspring from hypertensive families. Blood Press 9: 250-254. Jo¨nsson EG, Bah J, Melke J, Jamra RA, Schumacher J, et al. 2004. Monoamine related functional gene variants and relationships to monoamine metabolite concentrations in CSF of healthy volunteers. BMC Psychiatry 4: 4. Jonsson EG, Geijer T, Gyllander A, Terenius L, Sedvall GC. 1998. Failure to replicate an association between a rare allele of a tyrosine hydroxylase gene microsatellite and schizophrenia. Eur Arch Psychiatry Clin Neurosci 248: 61-63. Jonsson EG, Norton N, Forslund K, Mattila‐Evenden M, Rylander G, et al. 2003. Association between a promoter variant in the monoamine oxidase A gene and schizophrenia. Schizophr Res 61: 31-37. Jonsson E, Sedvall G, Brene S, Gustavsson JP, Geijer T, et al. 1996. Dopamine‐related genes and their relationships to monoamine metabolites in CSF. Biol Psychiatry 40: 1032-1043. Jorm A, Henderson A, Jacomb P, Christensen H, Korten A, et al. 2000. Association of a functional polymorphism of the monoamine oxidase A gene promoter with personality and psychiatric symptoms. Psychiatr Genet 10: 87-90. Kang J, Shi Y, Xiang B, Qu B, Su W, et al. 2005. A nuclear function of [beta]‐arrestin1 in GPCR signaling: Regulation of histone acetylation and gene transcription. Cell 123: 833-847. Karayiorgou M, Altemus M, Galke BL, Goldman D, Murphy DL, et al. 1997. Genotype determining low catechol‐O‐methyltransferase activity as a risk factor for obsessive‐compulsive disorder. Proc Natl Acad Sci USA 94: 4572-4575. Karayiorgou M, Gogos J. 1997. A turning point in schizophrenia genetics. Neuron 19: 967-979. Karayiorgou M, Gogos JA, Galke BL, Wolyniec PS, Nestadt G, et al. 1998. Identification of sequence variants and analysis of the role of the catechol‐O‐methyl‐transferase gene in schizophrenia susceptibility. Biolo Psychiatry 43: 425-431. Karayiorgou M, Morris MA, Morrow B, Shprintzen RJ, Goldberg R, et al. 1995. Schizophrenia susceptibility associated with interstitial deletions of chromosome 22q11. Proc Natl Acad Sci USA 92: 7612-7616. Kawada Y, Hattori M, Fukuda R, Arai H, Inoue R, et al. 1995. No evidence of linkage or association between tyrosine hydroxylase gene and affective disorder. J Affect Disord 34: 89-94. Kelsoe J, Ginns E, Egeland J, Gerhard D, Goldstein A, et al. 1989. Re‐evaluation of the linkage relationship between chromosome 11p loci and the gene for bipolar affective disorder in the old order amish. Nature 342: 238-243. Kirov G, Norton N, Jones I, McCandless F, Craddock N, et al. 1999. A functional polymorphism in the promoter of

143

144

6

Molecular genetics of brain noradrenergic neurotransmission

monoamine oxidase A gene and bipolar affective disorder. Int J Neuropsychopharmcol 2: 293-298. Klimek V, Stockmeier C, Overholser J, Meltzer HY, Kalka S, et al. 1997. Reduced levels of norepinephrine transporters in the locus coeruleus in major depression. J Neurosci 17: 8451-8458. Kohnke MD, Zabetian CP, Anderson GM, Kolb W, Gaertner I, et al. 2002. A genotype‐controlled analysis of plasma dopamine [beta]‐hydroxylase in healthy and alcoholic subjects: Evidence for alcohol‐related differences in noradrenergic function. Biol Psychiatry 52: 1151-1158. Korner J, Fritze J, Propping P. 1990. RFLP alleles at the tyrosine hydroxylase locus: No association found to affective disorders. Psychiatry Res 32: 275-280. Korner J, Rietschel M, Hunt N, Castle D, Gill M, et al. 1994. Association and haplotype analysis at the tyrosine hydroxylase locus in a combined German–British sample of manic depressive patients and controls. Psychiatr Genet 4: 167-175. Kunugi H, Ishida S, Kato T, Tatsumi M, Sakai T, et al. 1999. A functional polymorphism in the promoter region of monoamine oxidase‐A gene and mood disorders. Mol Psychiatry 4: 393-395. Kunugi H, Nanko S, Ueki A, Otsuka E, Hattori M, et al. 1997a. High and low activity alleles of catechol‐O‐methyltransferase gene: Ethnic difference and possible association with Parkinson’s disease. Neurosci Lett 221: 202-204. Kunugi H, Vallada HP, Hoda F, Kirov G, Gill M, et al. 1997b. No evidence for an association of affective disorders with high‐ or low‐activity allele of catechol‐O‐methyltransferase gene. Biol Psychiatry 42: 282-285. Kurumaji A, Kuroda T, Yamada K, Yoshikawa T, Toru M. 2001. An association of the polymorphic repeat of tetranucleotide (TCAT) in the first intron of the human tyrosine hydroxylase gene with schizophrenia in a Japanese sample. J Neural Transm 108: 489-495. Lachman HM, Kelsoe J, Moreno L, Katz S, Papolos DF. 1997. Lack of association of catechol‐O‐methyltransferase (COMT) functional polymorphism in bipolar affective disorder. Psychiatric Genet 7: 13-17. Lachman HM, Morrow B, Shprintzen R, Veit S, Parsia SS, et al. 1996b. Association of codon 108/158 catechol‐O‐ methyltransferase gene polymorphism with the psychiatric manifestations of velo‐cardio‐facial syndrome. Am J Med Genet 67: 468-472. Lachman HM, Nolan KA, Mohr P, Saito T, Volavka J. 1998. Association between catechol‐O‐methyltransferase genotype and violence in schizophrenia and schizoaffective disorder. Am J Psychiatry 155: 835-837. Lachman HM, Papolos DF, Saito T, Yu Y‐M, Szumlanski CL, et al. 1996a. Human catechol‐O‐methyltransferase pharmacogenetics: Description of a functional polymorphism and

its potential application to neuropsychiatric disorders. Pharmacogenetics 6: 243-250. Lan NC, Heinzmann C, Gal A, Klisak I, Orth U, et al. 1989. Human monoamine oxidase A and B genes map to Xp 11.23 and are deleted in a patient with norrie disease. Genomics 4: 552-559. Leboyer M, Malafosse A, Boularand S, Campion D, Gheysen F, et al. 1990. Tyrosine hydroxylase polymorphisms associated with manic‐depressive illness. Lancet 335: 1219. Leonard B. 2001. Stress, norepinephrine and depression. J Psychiatry Neurosci 26: Suppl: S11-S16. Lester HA, Cao Y, Mager S. 1996. Listening to neurotransmitter transporters. Neuron 17: 807-810. Li T, Vallada H, Curtis D, Arranz M, Xu K, et al. 1997. Catechol‐O‐methyltransferase Val158Met polymorphism: Frequency analysis in Han Chinese subjects and allelic association of the low activity allele with bipolar affective disorder. Pharmacogenetics 7: 349-353. Lim LC, Gurling H, Curtis D, Brynjolfsson J, Petursson H, et al. 1993. Linkage between tyrosine hydroxylase gene and affective disorder cannot be excluded in two of six pedigrees. Am J Med Genet 48: 223-228. Lobos EA, Todd RD. 1997. Cladistic analysis of disease association with tyrosine hydroxylase: Application to manic‐ depressive disease and alcoholism. Am J Med Genet 74: 289-295. Lohmueller KE, Pearce CL, Pike M, Lander ES, Hirschhorn JN. 2003. Meta‐analysis of genetic association studies supports a contribution of common variants to susceptibility to common disease. Nat Genet 33: 177-182. Lotta T, Vidgren J, Tilgmann C, Ulmanen I, Melen K, et al. 1995. Kinetics of human soluble and membrane‐bound catechol‐O‐methyltransferase: A revised mechanism and description of the thermolabile variant of the enzyme. Biochemistry 34: 4202-4210. Lundstrom K, Tenhunen J, Tilgmann C, Karhunen T, Panula P, et al. 1995. Cloning, expression and structure of catechol‐ O‐methyltransferase. Biochim Biophys Acta Protein Structure and Molecular Enzymology 1251: 1-10. Malafosse A, Leboyer M, d’Amato T, Amadeo S, Abbar M, et al. 1997. Manic depressive illness and tyrosine hydroxylase gene: Linkage heterogeneity and association. Neurobiol Dis 4: 337-349. Mallet J. 1996. The TiPS/TINS lecture. Catecholamines: From gene regulation to neuropsychiatric disorders. Trends Pharmacol Sci 17: 129-135. Mandela P, Ordway GA. 2006. The norepinephrine transporter and its regulation. J Neurochem 97: 310-333. Masson J, Sagne C, Hamon M, Mestikawy SE. 1999. Neurotransmitter transporters in the central nervous system. Pharmacol Rev 51: 439-464.

Molecular genetics of brain noradrenergic neurotransmission Mattay VS, Goldberg TE., Fera F, Hariri AR., Tessitore A, et al. 2003. Catechol‐O‐methyltransferase val158‐met genotype and individual variation in the brain response to amphetamine. Proc Natl Acad Sci USA 100: 6186-6191. Melia K, Duman R. 1991. Involvement of corticotropin‐ releasing factor in chronic stress regulation of the brain noradrenergic system. Proc Natl Acad Sci USA 88: 8382-8386. Melia K, Rasmussen K, Terwilliger R, Haycock J, Nestler E, et al. 1992. Coordinate regulation of the cyclic AMP system with firing rate and expression of tyrosine hydroxylase in the rat locus coeruleus: Effects of chronic stress and drug treatments. J Neurochem 58: 494-502. Meloni R, Albanese V, Ravassard P, Treilhou F, Mallet J. 1998. A tetranucleotide polymorphic microsatellite, located in the first intron of the tyrosine hydroxylase gene, acts as a transcription regulatory element in vitro. Hum Mol Genet 7: 423-428. Meloni R, Laurent C, Campion D, Ben Hadjali B, Thibaut F, et al. 1995b. A rare allele of a microsatellite located in the tyrosine hydroxylase gene found in schizophrenic patients. C R Acad Sci III 318: 803-809. Meloni R, Leboyer M, Bellivier F, Barbe B, Samolyk D, et al. 1995a. Association of manic‐depressive illness with tyrosine hydroxylase microsatellite marker. Lancet 345: 932. Meyer E, Wiegand P, Rand SP, Kuhlmann D, Brack M, et al. 1995. Microsatellite polymorphisms reveal phylogenetic relationships in primates. J Mol Evol 41: 10-14. Moore H, West AR, Grace AA. 1999. The regulation of forebrain dopamine transmission: Relevance to the pathophysiology and psychopathology of schizophrenia. Biol Psychiatry 46: 40-55. Morilak DA, Barrera G, Echevarria DJ, Garcia AS, Hernandez A, et al. 2005. Role of brain norepinephrine in the behavioral response to stress. Prog Neuropsychopharmacol Biol Psychiatry—Experimental Stress: From Basic to Clinical Aspects 29: 1214-1224. Mu¨ller Smith K, Daly M, Fischer M, Yiannoutsos CT, Bauer L, et al. 2003. Association of the dopamine beta hydroxylase gene with attention deficit hyperactivity disorder: Genetic analysis of the Milwaukee longitudinal study. Am J Med Genet 119B: 77-85. Mynett‐Johnson LA, Murphy VE, Claffey E, Shields DC, McKeon P. 1998. Preliminary evidence of an association between bipolar disorder in females and the catechol‐O‐ methyltransferase gene. Psychiatr Genet 8: 221-225. Neer EJ. 1995. Heterotrimeric C proteins: Organizers of transmembrane signals. Cell 80: 249-257. Nelson JC. 1999. A review of the efficacy of serotonergic and noradrenergic reuptake inhibitors for treatment of major depression. Biol Psychiatry 46: 1301-1308. Njus D, Kelley P, Harnadek G. 1986. Bioenergetics of secretory vesicles. Biochim Biophys Acta 853: 237-265.

6

Ohmori O, Shinkai T, Kojima H, Terao T, Suzuki T, et al. 1998. Association study of a functional catechol‐O‐methyltransferase gene polymorphism in Japanese schizophrenics. Neurosci Lett 243: 109-112. Ordway G. 1997. Pathophysiology of the locus coeruleus in suicide. Ann N Y Acad Sci 836: 233-252. Pakstis A, Kidd J, Castiglione C, Kidd K. 1991. Status of the search for a major genetic locus for affective disorder in the old order amish. Hum Genet 87: 475-483. Palmatier MA, Kang AM, Kidd KK. 1999. Global variation in the frequencies of functionally different catechol‐O‐ methyltransferase alleles. Biol Psychiatry 46: 557-567. Perez de Castro I, Santos J, Torres P, Visedo G, Saiz‐Ruiz J, et al. 1995. A weak association between TH and DRD2 genes and bipolar affective disorder in a Spanish sample. J Med Genet 32: 131-134. Persson ML, Wasserman D, Jonsson EG, Bergman H, Terenius L, et al. 2000. Search for the influence of the tyrosine hydroxylase (TCAT)(n) repeat polymorphism on personality traits. Psychiatry Res 95: 1-8. Polymeropoulos M, Xiao H, Rath D, Merril C. 1991. Tetranucleotide repeat polymorphism at the human tyrosine hydroxylase gene (TH). Nucleic Acids Res 19: 3753. de Pontual L, Nepote V, Attie‐Bitach T, Al Halabiah H, Trang H, et al. 2003. Noradrenergic neuronal development is impaired by mutation of the proneural HASH‐1 gene in congenital central hypoventilation syndrome (ondine’s curse). Hum Mol Genet 12: 3173-3180. Puers C, Hammond HA, Jin L, Caskey T, Schumm JW. 1993. Identification of repeat sequence heterogeneity at the polymorphic short tandem repeat locus HUMTH01 (AATG)n and reassignment of alleles in population analysis by using a locus‐specific allelic ladder. Am J Hum Genet 53: 953-958. Pulver AE, Karayiorgou M, Lasseter VK, Wolyniec P, Kasch L, et al. 1994b. Follow‐up of a report of a potential linkage for schizophrenia on chromosome 22q12–q13.1: Part 2. Am J Med Genet 54: 44-50. Pulver AE, Karayiorgou M, Wolyniec PS, Lasseter VK, Kasch L, et al. 1994a. Sequential strategy to identify a susceptibility gene for schizophrenia: Report of potential linkage on chromosome 22q12–q13.1: Part 1. Am J Med Genet 54: 36-43. Sabban EL, Kvetnansky R. 2001. Stress‐triggered activation of gene expression in catecholaminergic systems: Dynamics of transcriptional events. Trends Neurosci 24: 91-98. Sabol S, Hu S, Hamer D. 1998. A functional polymorphism in the monoamine oxidase A gene promoter. Hum Genet 103: 273-279. Sander T, Harms H, Rommelspacher H, Hoehe M, Schmidt LG. 1998. Possible allelic association of a tyrosine hydroxylase polymorphism with vulnerability to alcohol‐ withdrawal delirium. Psychiatr Genet 8: 13-17.

145

146

6

Molecular genetics of brain noradrenergic neurotransmission

Sassone‐Corsi P, Lamph WW, Kamps M, Verma IM. 1988. Fos‐associated cellular p39 is related to nuclear transcription factor AP‐1. Cell 54: 553-560. Schatzberg A. 2000. Clinical efficacy of reboxetine in major depression. J Clin Psychiatry 61: 31-38. Schildkraut JJ. 1965. The catecholamine hypothesis of affective disorders: A review of supporting evidence. Am J Psychiatry 122: 509-522. Schultz, W, Dayan P, Montague P. 1997. A neural substrate of prediction and reward. Science 275: 1593-1599. Schulze T, Muller D, Krauss H, Scherk H, Ohlraun S, et al. 2000. Association between a functional polymorphism in the monoamine oxidase A gene promoter and major depressive disorder. Am J Med Genet 96: 801-803. SerrettiA, Macciardi F, Cusin C, Verga M, Pedrini S, 1998a. Tyrosine hydroxylase gene in linkage disequilibrium with mood disorders. Mol Psychiatry 3: 169-174. Serretti A, Macciardi F, Verga M, Cusin C, Pedrini S, et al. 1998b. Tyrosine hydroxylase gene associated with depressive symptomatology in mood disorder. Am J Med Genet 81: 127-130. Servan‐Schreiber D, Printz H, Cohen J. 1990. A network model of catecholamine effects: Gain, signal‐to‐noise ratio, and behavior. Science 249: 892-895. Shifman S, Bronstein M, Sternfeld M, Pisante´ A, Weizman A, et al. 2004. COMT: A common susceptibility gene in bipolar disorder and schizophrenia. Am J Med Genet 128B: 61-64. Shifman S, Bronstein M, Sternfeld M, Pisante‐Shalom A, Lev‐Lehman E, et al. 2003. A highly significant association between a COMT haplotype and schizophrenia. Am J Hum Genet 71: 1296-1302. Shih JC, Chen K, Ridd, J. 1999. Monoamine oxidase: From genes to behavior. Annu Rev Neurosci 22: 197-217. Smyth C, Kalsi G, Brynjolfsson J, O’Neill J, Curtis D, et al. 1996. Further tests for linkage of bipolar affective disorder to the tyrosine hydroxylase gene locus on chromosome 11p15 in a new series of multiplex British affective disorder pedigrees. Am J Psychiatry 153: 271-274. Souery D, Lipp O, Mahieu B, Mendelbaum K, De Martelaer V, et al. 1996. Association study of bipolar disorder with candidate genes involved in catecholamine neurotransmission: DRD2, DRD3, DAT1, and TH genes. Am J Med Genet 67: 551-555. Souery D, Lipp O, Rivelli SK, Massat I, Serretti A, et al. 1999. Tyrosine hydroxylase polymorphism and phenotypic heterogeneity in bipolar affective disorder: A multicenter association study. Am J Med Genet 88: 527-532. Southwick S, Yehuda R, Giller EJ. 1993. Personality disorders in treatment‐seeking combat veterans with posttraumatic stress disorder. Am J Psychiatry 150: 1020-1023. Starke K. 2001. Presynaptic autoreceptors in the third decade: Focus on alpha2‐adrenoceptors. J Neurochem 78: 685-693.

Stolk JM, Hurst JH, Nisula BC. 1982. Regulation and inheritance of dopamine‐beta‐hydroxylase. Behav Genet 12: 37-52. Strous RD, Bark N, Woerner M, Lachman HM. 1997. Lack of association of a functional catechol‐O‐methyltransferase gene polymorphism in schizophrenia. Biol Psychiatry 41: 493-495. Sugama S, Namihira T, Matsuoka R, Taira N, Eto Y, et al. 1999. Psychiatric inpatients and chromosome deletions within 22q11.2. J Neurol Neurosurg Psychiatry 67: 803-806. Sullivan G, Coplan J, Kent J, Gorman J. 1999. The noradrenergic system in pathological anxiety: A focus on panic with relevance to generalized anxiety and phobias. Biol Psychiatry 46: 1205-1218. Syagailo Y, Stober G, Grassle M, Reimer E, Knapp M, et al. 2001. Association analysis of the functional monoamine oxidase a gene promoter polymorphism in psychiatric disorders. Am J Med Genet 105: 168-171. Syvanen A‐C, Tilgmann C, Rinne J, Ulmanen I. 1997. Genetic polymorphism of catechol‐O‐methyltransferase (COMT): Correlation of genotype with individual variation of S‐COMT activity and comparison of the allele frequencies in the normal population and Parkinsonian patients in Finland. Pharmacogenetics 7: 65-71. Taussig R, Gilman AG. 1995. Mammalian membrane‐bound adenylyl cyclases. J Biol Chem 270: 1-4. Tenhunen J, Salminen M, Lundstrom K, Savolainen R, Ulmanen I. 1994. Genomic organization of the human catechol‐O‐methyltransferase gene and its expression from two distinct promoters. Eur J Biochem 223: 1049-1059. Thibaut F, Ribeyre JM, Dourmap N, Meloni R, Laurent C, et al. 1997. Association of DNA polymorphism in the first intron of the tyrosine hydroxylase gene with disturbances of the catecholaminergic system in schizophrenia. Schizophr Res 23: 259-264. Turecki G, Rouleau GA, Mari J, Joober R, Morgan K. 1997. Lack of association between bipolar disorder and tyrosine hydroxylase: A meta‐analysis. Am J Med Genet 74: 348-352. Vandenbergh DJ, Rodriguez LA, Miller IT, Uhl GR, Lachman HM. 1997. High‐activity catechol‐O‐methyltransferase allele is more prevalent in polysubstance abusers. Am J Med Genet—Neuropsychiatric Genetics 74: 439-442. Versiani M, Cassano G, Perugi G, Benedetti A, Mastalli L, et al. 2002. Reboxetine, a selective norepinephrine reuptake inhibitor, is an effective and well‐tolerated treatment for panic disorder. J Clin Psychiatry 63: 31-37. Wei J, Hemmings GP. 1999. Lack of evidence for association between the COMT locus and schizophrenia. Psychiatr Genet 9: 183-186. Wei J, Ramchand CN, Hemmings GP. 1995. Association of polymorphic VNTR region in the first intron of the human

Molecular genetics of brain noradrenergic neurotransmission TH gene with disturbances of the catecholamine pathway in schizophrenia. Psychiatr Genet 5: 83-88. Wei J, Ramchand CN, Hemmings GP. 1997. Possible association of catecholamine turnover with the polymorphic (TCAT)n repeat in the first intron of the human tyrosine hydroxylase gene. Life Sci 61: 1341-1347. Wei J, Ramchand CN, Hemmings G. 1998. TaqI polymorphic sites at the human dopamine beta‐hydroxylase gene possibly associated with biochemical alterations of the catecholamine pathway in schizophrenia. Psychiatr Genet 8: 19-24. Weinberger DR, Egan MF, Bertolino A, Callicott JH, Mattay VS, et al. 2001. Prefrontal neurons and the genetics of schizophrenia. Biol Psychiatry 50: 825-844. Weinshilboum R. 1978. Human biochemical genetics of plasma dopamine‐[beta]‐hydroxylase and erythrocyte catechol‐ O‐methyltransferase. Hum Genet 45: 101-112. Wigg K, Zai G, Schachar R, Tannock R, Roberts W, et al. 2002. Attention deficit hyperactivity disorder and the gene for dopamine beta‐hydroxylase. Am J Psychiatry 159: 1046-1048. Woodward D, Moises H, Waterhouse B, Yeh H, Cheun J. 1991. Modulatory actions of norepinephrine on neural circuits. Adv Exp Med Biol 287: 193-208. Xie T, Ho SL, Li LSW, Ma OCK. 1997. G/A1947 polymorphism in catechol‐O‐methyltransferase (COMT) gene in Parkinson’s disease. Mov Disord 12: 426-427. Yamamoto K, Cubells J, Gelernter J, Benkelfat C, Lalonde P, et al. 2003. Dopamine beta‐hydroxylase (DBH) gene and schizophrenia phenotypic variability: A genetic association study. Am J Med Genet 117B: 33-38.

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Yamamoto K, Hornykiewicz O. 2004. Proposal for a noradrenaline hypothesis of schizophrenia. Prog Neuropsychopharmacol Biol Psychiatry 28: 913-922. Young LJ, Nilsen R, Waymire KG, Mac Gregor GR, Insel TR. 1999. Increased affiliative response to vasopressin in mice expressing the V1a receptor from a monogamous vole. Nature 400: 766-768. Zabetian CP, Anderson GM, Buxbaum SG, Elston RC, Ichinose H, et al. 2001. A quantitative‐trait analysis of human plasma‐dopamine beta‐hydroxylase activity: Evidence for a major functional polymorphism at the DBH locus. Am J Hum Genet 68: 515-522. Zabetian CP, Buxbaum SG, Elston RC, Ko¨hnke MD, Anderson GM, et al. 2003. The structure of linkage disequilibrium at the DBH locus strongly influences the magnitude of association between diallelic markers and plasma dopamine—hydroxylase activity. Am J Hum Genet 72: 1389-1400. Zhang L, Rao F, Wessel J, Kennedy BP, Rana BK, et al. 2004. Functional allelic heterogeneity and pleiotropy of a repeat polymorphism in tyrosine hydroxylase: Prediction of catecholamines and response to stress in twins. Physiol Genomics 19: 277-291. Zhu MY, Klimek V, Dilley GE, Haycock JW, Stockmeier C, et al. 1999. Elevated levels of tyrosine hydroxylase in the locus coeruleus in major depression. Biol Psychiatry 46: 1275-1286. Zhuma T, Tyrrell R, Sekkali B, Skavdis G, Saveliev A, et al. 1999. Human HMG box transcription factor HBP1: A role in hCD2 LCR function. EMBO J 18: 6396-6406.

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L. G. Harsing Jr.

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 151

2 2.1 2.2 2.3 2.4

The Synthetic Pathway of Dopamine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 152 Tyrosine Hydroxylase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 152 Estimation of Dopamine Turnover Rate from Dopamine Synthesis Inhibition . . . . . . . . . . . . . . . . . . 154 L‐Aromatic Amino Acid Carboxylase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 154 Estimation of Dopamine Synthesis Rate from Dopa Decarboxylase Inhibition . . . . . . . . . . . . . . . . . . 154

3 3.1 3.2 3.3

The Degradative Pathway of Dopamine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 155 Monoamine Oxidase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 155 Catechol‐O‐Methyl‐Transferase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 155 Estimation of Dopamine Turnover Rate by Calculation of Metabolites/Dopamine Ratio . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 155

4 Storage of Dopamine in Neuronal Pools . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 156 4.1 Vesicular Monoamine Transporter . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 158 5 5.1 5.2 5.3

Dopamine Plasma Membrane Transporter . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 158 Structure of Dopamine Transporter . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 158 Operation of Dopamine Transporter . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 159 Drugs Acting on Dopamine Transporter . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 159

6 6.1 6.2 6.3 6.4 6.5

Dopamine Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 159 Classification of Dopamine Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 160 Structure of Dopamine Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 160 Postsynaptic Dopamine Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 161 Presynaptic Dopamine Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 161 Regional Distribution of Dopamine Autoreceptors Assessed by the g‐Butyrolactone Model . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 162 6.6 Changes in Dopamine Receptor Sensitivity and Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 162 7 7.1 7.2 7.3 7.4

Dopamine Release from Neuronal Stores . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 163 Action Potential Propagation‐Induced Dopamine Release . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 163 Dopamine Release Evoked by Reverse‐Mode Operation of Dopamine Transporters . . . . . . . . . . . . . 164 Dopamine Release Evoked by Ion Channel‐Coupled Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 165 Regulation of Dopamine Release . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 165

8 Dopaminergic Innervations in the Central Nervous System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 165 8.1 Dopamine in the Striatum . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 166

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8.2 Dopamine in the Cerebral Cortex . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 166 9 Altered Dopaminergic Neurotransmission . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 166 9.1 Neurotoxins Used for Destruction of Dopaminergic Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 166 9.2 Dopaminergic Neurotransmission in Knockout Mice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 167 10

Conclusions and Future Avenues . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 167

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Abstract: The great majority of dopaminergic neurons in the brain (in human 300,000–400,000 cells) is organized in three nuclei, the substantia nigra pars compacta, the ventral tegmental area and the arcuate nucleus. Long pathways originating from these areas project to the neostriatum, the cerebral cortex and limbic structures, and the hypothalamus. Dopaminergic neurotransmission is believed to have a central role in the integration of the reward system; it organizes complete motor programs and voluntary movements, and regulates some endocrine hormone secretion. In pathological conditions, this neurotransmitter system mediates extrapyramidal side effects, drug abuse and dependence, and it is responsible to develop psychiatric disorders such as schizophrenia and attention‐deficit hyperactivity disorder. There are two enzymes, tyrosine hydroxylase and dopa decarboxylase, involved in the synthesis and two enzymes, monoamine oxidase and catechol‐O‐ methyl transferase, take part in the degradation of dopamine. Dopamine in the axon terminals is stored in the cytoplasm and in at least two different vesicular pools. Vesicular monoamine transporter concentrates dopamine into vesicles; this storage protects dopamine from enzymatic degradation, retards dopamine from diffusion out from nerve endings, and serves as pool from where dopamine can be released in response to physiological stimuli. Dopamine release from axon terminals may occur with different mechanisms: action potential propagation‐induced membrane depolarization evokes release from the vesicular stores, whereas the amphetamine class of drugs may induce dopamine efflux by forcing dopamine transporter to operate in reverse mode. The action of dopamine released is primarily terminated by its reuptake into the presynaptic terminals. Dopamine transporter is a transmembrane protein that removes dopamine from the synaptic cleft before it can escape into the biophase. Dopamine transporter knockout mice exhibit a wide range of deficit in dopaminergic functions and altered dopamine‐mediated behavior. There is growing evidence for the importance of extracellular dopamine in the regulation of nonsynaptic neurotransmission, drug actions, and mediation of some dopamine‐related psychiatric disorders. Dopamine, if once released into the synaptic cleft, acts on dopamine receptors. Five dopamine receptors, the D1‐like D1 and D5 and the D2‐like D2, D3, and D4 receptors were identified. Dopamine receptors are coupled to G proteins and they positively or negatively regulate intracellular messenger cascade in which the phosphoprotein DARP‐32 possesses central role. Based upon their expression, presynaptic and postsynaptic dopamine receptors can be distinguished. Response to postsynaptic D1 and D2 receptors alters cellular messenger cascades and that to presynaptic D2 receptors stimulation influences dopamine release and synthesis and neuronal firing rate. Newly synthesized compounds with affinity to dopamine D2/D3 receptors or inhibitors of dopamine degradation are now clinically tested for treatment of schizophrenia and Parkinson’s disease. List of Abbreviations: AADC, amino acid carboxylase; AC, adenylate cyclase; cAMP, cyclic adenosine 50 ‐monophosphate; COMT, catechol‐O‐methyl‐transferase; DA, dopamine; DARPP‐32, dopamine‐ and cAMP‐regulated phosphoprotein‐32; DAT, dopamine plasma membrane transporter; DOPAC, 3,4‐dihydroxyphenylacetic acid; DOPA, dihydroxyphenylalanine; GABA, g‐amino‐butyric acid; GBL, g‐butyrolactone; G‐protein, guanine nucleotide binding protein; HVA, homovanillic acid; MAO, monoamine oxidase; MPPþ, 1‐methyl‐4‐phenylpyridinium; MPTP, 1‐methyl‐4‐phenyl‐1,2,3,6‐tetrahydropyridine; NO, nitric oxide; NOS, nitric oxide synthase; pDARPP‐32, phosphorylated dopamine‐ and cAMP‐regulated phosphoprotein‐32; PKA, protein kinase A; PP‐1, protein phosphatase‐1; PP‐2B, protein phosphatase‐2B; SAM, S‐adenosyl‐methionine; SLC, solute carrier; TH, tyrosine hydroxylase; 3‐MT, 3‐methoxytyramine; TR, turnover rate; VMATs, vesicular monoamine transporter proteins

1

Introduction

Dopamine (3,4‐dihydroxyphenylalanine) is an endogenous compound containing a benzene ring with two hydroxyl substituents (catechol nucleus) and an aminoethyl group attached to the substituted ring. Dopamine and other compounds with similar molecular structures (norepinephrine and epinephrine) belong to catecholamines and they are often referred in neurosciences as biogenic amines or monoamine neurotransmitters. Dopamine is an important neurotransmitter in the central nervous system and mediates a number of physiological regulations and also involved in the development of neurological and psychiatric disorders. Thus, dopamine has a major role in the pathology of control of movement (Parkinson’s disease)

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or in psychiatric disorders such as schizophrenia and attention deficit hyperactivity disorder. In addition, dopaminergic neurotransmission has a central role in the mechanism of drug abuse associated with dependence and also in the integration of the reward system of the brain. Long dopaminergic projections in the central nervous system that relate to these functions were visualized first by a fluorescent method described by Dahlstrom and Fuxe (1964). Dopamine and other biogenic amines (norepinephrine, epinephrine, serotonin, and histamine) are members of the classic neurotransmitters. There are several differences between the morphological appearance and neurochemistry of the classic and nonclassic (peptide) neurotransmitters. Storage vesicles, when they are present, are smaller for classic neurotransmitters and they are larger for nonclassic neurotransmitters. Another different feature is that classic neurotransmitters or, in some cases, their metabolic products are subject to reuptake but there is no energy dependent, high‐affinity transport system for peptide neurotransmitters. Moreover, classic neurotransmitters are synthesized in the presynaptic axon terminals, the synthesis of nonclassic transmitters occurs in the cell body and the precursor proteins reach nerve endings by axonal transport.

2

The Synthetic Pathway of Dopamine

There are two enzymatic steps involved in dopamine synthesis (Von Bohlen und Halbach and Dermietzel, 2002). The biosynthetic pathway of dopamine begins with the amino acid precursor tyrosine (> Table 7-1). Phenylalanine is converted to tyrosine by the enzyme phenylalanine hydroxylase although dopamine biosynthesis is usually considered to begin with tyrosine (> Figure 7-1). Tyrosine is hydroxylated at

. Table 7-1 Enzymes involved in the biosynthesis and degradation of dopamine Enzymes Tyrosine hydroxylase L‐aromatic amino acid carboxylase Dopamine‐b‐hydroxylase

Substrate Tyrosine Dopa

Product Dihydroxy‐phenylalanine Dopamine

Dopamine

Norepinephrine

Monoamine oxidase A

Norepinephrine

Monoamine oxidase B

Dopaminea

Catechol‐O‐methyl‐ transferase

Dopamine Norepinephrine

3,4‐Dihydroxy‐ phenylglycolaldehyde 3,4‐Dihydroxyphenyl‐ acetaldehyde 3‐Methoxytyramine Normetanephrine

Inhibitors a‐Methyl p‐tyrosine Carbidopa Benserazide Copper chalators FLA‐63 Clorgyline L‐deprenyl Tolcapone

a

Dopamine is a substrate for type A monoamine oxidase in rat striatum (Demarest et al., 1980), but it is monoamine oxidase B substrate in human (Glover et al., 1977)

position 3 by the enzyme tyrosine hydroxylase (TH) and 3,4‐dihydroxy‐L‐phenylalanine (L‐dopa) is formed. Dopamine synthesis is followed by the next step, when L‐aromatic amino acid decarboxylase converts dopa to dopamine (Deutch and Roth, 2004).

2.1 Tyrosine Hydroxylase Tyrosine hydroxylase catalyzes the addition of hydroxyl group to the meta position of tyrosine, thus forming L‐dopa. Physiological tyrosine concentrations saturate tyrosine hydroxylase and increase of tyrosine concentrations usually does not elevate the rate of dopamine synthesis. Tyrosine hydroxylase is the

Dopamine and the dopaminergic systems of the brain

7

. Figure 7-1 The life cycle of dopamine

rate‐limiting step in dopamine synthesis and the enzyme controls the neuronal concentrations of dopamine. Tyrosine hydroxylase is a mixed‐function oxidase that has moderate substrate specificity; it hydroxylates not only tyrosine but phenylalanine also, particularly in conditions when phenylalanine hydroxylase in suppressed (phenylketonuria). The actions of tyrosine hydroxylase require tetrahydrobiopterin cofactor as hydrogen donor, Fe2þ ion, and molecular oxygen (Thony et al., 2000). Activation of dopaminergic neurons leads to increase of tyrosine hydroxylase activity and dopaminergic neurotransmission is elevated. The short‐term regulation of tyrosine hydroxylase occurs through the posttranslational levels whereas long‐term regulation of tyrosine hydroxylase activity can occur through transcriptional regulation of the gene. Rapid and short term activation of tyrosine hydroxylase occur through phosphorylation and dephosphorylation of at least four serine residues (Ser‐8, Ser‐19, Ser‐31, Ser‐40) in the N terminal part of the enzyme by a series of distinct protein kinases including protein kinase A, protein kinase C, and Ca2þ/calmodulin‐dependent protein kinase II. The conformational changes of the enzyme during phosphorylation result in higher affinity to tetrahydrobiopterin cofactor and lower affinity to dopamine and thus, the endproduct inhibition of the enzyme is decreasing. Tyrosine hydroxylase expression can be either upregulated or downregulated by different drugs such as nicotine, caffeine, morphine, or antidepressants by activating or repressing transcriptional regulatory elements of tyrosine hydroxylase gene promoter. These regulatory elements may include cAMP response element (CRE), glycocorticoid response element (GRE), activator proteins‐1 (AP‐1) or NF‐kB sites (Nestler et al., 2001).

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2.2 Estimation of Dopamine Turnover Rate from Dopamine Synthesis Inhibition Analogs of tyrosine (a‐methyl‐p‐tyrosine, 3‐iodotyrosine) are competitive inhibitor of tyrosine hydroxylase (Walter et al., 1984; Snyder et al., 1990). a‐Methyl‐p‐tyrosine is widely used to study catecholamine neurotransmission functions, including dopamine turnover rate (Walter et al., 1984; Vizi et al., 1986). Dopamine biosynthesis can be assessed by estimating in vivo activity of tyrosine hydroxylase after treatment with a‐methyl‐p‐tyrosine. Apparent rate of dopamine turnover in brain regions can be determined by following the rate of decline of dopamine concentrations at certain time intervals after i.p. injection of a‐methyl‐p‐tyrosine. The levels of dopamine (as well as its precursors and metabolites) can be determined by HPLC‐electrochemistry. Drugs that are tested to alter dopamine turnover, are usually given before a‐methyl‐p‐tyrosine administration. After blockade of synthesis by a‐methyl‐p‐tyrosine, dopamine concentrations decline at a rate that is proportional to concentration, i.e., log½DA
t ¼ log½DA
0  0:434 kt where [DA]0 is the initial level, [DA]t is the level at time t, and k is the rate constant of dopamine efflux. The turnover rate is product of steady‐state level and of k (the rate constant of dopamine decline): TRDA ¼ k[DA]0. The turnover time of dopamine (the time required to replace the amine pool) is then calculated as Tt ¼1/k. The validity of this method for determining turnover rates of dopamine depends on a‐methyl‐p‐ tyrosine being maintained at an inhibitory level. This was evidenced in that the complete blockade of the enzyme resulted in a decline of dopamine level to an almost zero value at an exponential rate and by the fact that higher doses of a‐methyl‐p‐tyrosine does not increase further the decline.

2.3 L‐Aromatic Amino Acid Carboxylase The hydroxylation of tyrosine by tyrosine hydroxylase generates L‐dopa, which is then decarboxylated by the enzyme L‐aromatic amino acid carboxylase (AADC), also referred as dopa decarboxylase. This enzyme has low Km and high Vmax values and levels of L‐dopa are virtually unmeasurable in the brain under basal conditions. This is because the activity of dopa decarboxylase is so high that L‐dopa is converted into dopamine almost instantaneously. L‐aromatic AADC has low substrate specificity and decarboxylases tyrosine and tryptophane as well as other aromatic amino acids. L‐aromatic AADC is a cytoplasmic enzyme that is present in catecholamine‐ and serotonin‐containing neurons. The enzyme requires pyridoxal 5‐phosphate as a cofactor for its activity. L‐aromatic AADC is exploited in the treatment of Parkinson’s disease by giving L‐dopa to patients to enhance dopamine production in the remaining dopaminergic axon terminals. As dopamine does not cross the blood–brain barrier and L‐dopa readily enters the brain, the latter is used for substitution therapy. A series of L‐dopa analogs such as a‐methyldopa and carbidopa inhibit dopa decarboxylase. Benserazide and RO4‐4602 are also known inhibitors of the enzyme. NSD‐1015 (3‐hydroxybenzylhydrazine) is a centrally active decarboxylase inhibitor and it is widely used to study drugs modifying dopamine synthesis.

2.4 Estimation of Dopamine Synthesis Rate from Dopa Decarboxylase Inhibition The synthesis rate of dopamine can be determined from measuring L‐dopa accumulation in brain tissue samples after inhibition of L‐aromatic amino acid decarboxylase with NSD‐1015. The accumulation of L‐dopa in the rat brain after administration of dopa decarboxylase inhibitors can be used as a measure to estimate tyrosine hydroxylase activity in vivo. L‐dopa accumulation is linear for a certain period after the

Dopamine and the dopaminergic systems of the brain

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administration of NSD‐1015. The increase in L‐dopa accumulation can be reduced by the dopamine D2 receptor agonist apomorphine and reversed by dopamine antagonists. NSD‐1015, when blocks L‐aromatic amino acid decarboxylase, also inhibits the conversion of the amino acid b‐phenylalanine into b‐phenylethylamine. Administration of NSD‐1015 also leads to elevation of 5‐hydroxytryptophan levels; a technique allows the measurement of serotonin synthesis rate in brain tissue samples.

3

The Degradative Pathway of Dopamine

3.1 Monoamine Oxidase Two major enzymes take part in dopamine catabolism: monoamine oxidase (MAO) and catechol‐O‐ methyl‐transferase (COMT) (Von Bohlen und Halbach and Dermietzel, 2002). MAO occurs in the tissues in the form of two isoenzymes, MAOA and MAOB. MAOA displays high affinity to norepinephrine and serotonin, MAOB exhibits the highest affinity to phenylethylamine (> Table 7-1). MAOA and MAOB have similar affinity to dopamine. MAOs oxidatively deaminate dopamine and its O‐methylated derivative, 3‐methoxytyramine (3‐MT), to form inactive and unstable aldehyde derivatives, 3,4‐dihydroxyphenylacetaldehyde and 3‐methoxy‐4‐hydroxyphenylacetaldehyde (> Figure 7-1). These aldehydes can then be further catabolized by aldehyde dehydrogenases to form corresponding acid metabolites, 3,4‐dihydroxyphenylacetic acid (DOPAC) and homovanillic acid (HVA). MAO requires flavin adenine dinucleotide as cofactor for its activity. The two different forms of MAO are derived from distinct genes and differ not only in their substrate specificity but also in their cellular locations and regulation by pharmacological agents. MAOA is selectively inhibited by clorgyline, and L‐deprenyl selectively inhibits MAOB. Both A and B forms of MAO are associated with the outer membrane of mitochondria. MAOB is also present in glial cells (Nestler et al., 2001).

3.2 Catechol‐O‐Methyl‐Transferase COMT acts to methylate catecholamines and requires S‐adenosyl‐methionine (SAM) as methyl donor for its activity. COMT is relatively nonspecific enzyme that transfers methyl groups from SAM to the meta‐ hydroxy group of catechols. This enzyme was identified in the synaptic cleft. COMT inhibitors (tolcapone, entacapone) increase dopamine levels within the synapses and prolong dopamine receptor activation.

3.3 Estimation of Dopamine Turnover Rate by Calculation of Metabolites/Dopamine Ratio The ratio of DOPAC to dopamine indicates the rate of dopamine metabolism, whereas changes in the levels of dopamine metabolites, DOPAC and HVA, reflect changes in MAO activity. Dopaminergic neuronal activity can be further estimated by calculation of DOPACþHVA/dopamine ratio. This ratio, which indicates alterations in the rate of dopamine turnover, was found to be changed in a number of pathological conditions (experimental parkinsonism or following stroke) as well as during drug treatments (Ogawa et al., 2000; Megyeri et al., 2007). Changes in the levels of 3‐methoxytyramine, a minor metabolite of dopamine, also reflect the turnover and utilization of dopamine. a‐Methyl‐p‐tyrosine produces a parallel decrease in dopamine and 3‐methoxytyramine levels in the striatum and nucleus accumbens. An enhanced 3‐methoxytyramine accumulation can be observed in rats pretreated with MAO inhibitors such as tranylcypromine or pargyline. In addition, accumulation of 3‐methoxytyramine in rat brain provides a sensitive assay to distinguish between dopamine‐releasing agents and uptake inhibitors (Heal et al., 1990).

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Storage of Dopamine in Neuronal Pools

Dopamine is synthesized in the nerve terminal cytoplasm to where enzymes involved in this process are transported from the cell body area (> Figure 7-2). Dopamine is then packed in storage vehicles by means of vesicular monoamine transporter (VMAT) proteins. Dopamine storage vesicles are present at high density within nerve terminals. Vesicles protect dopamine from inactivation by intraneuronal enzymes,

. Figure 7-2 The dopaminergic synapse: schematic diagram of the presynaptic nerve terminal, the synaptic cleft and the postsynaptic cell. At the presynaptic nerve terminals, dopamine is released by exocytosis or by reverse‐mode operation of the dopamine transporter. The released dopamine activates presynaptic D2 autoreceptors inhibiting further release of the neurotransmitter. This inhibition may be the consequence of inhibition of voltage‐ sensitive Ca2þ‐channels and/or inhibition of Kþpermeability. In addition, activation of D2 receptor‐coupled Gi/o protein leads to decreased cAMP production and inhibition of protein kinase A, which regulates proteins involved in the release process. For reversal of dopamine transporter, dopamine releaser drugs enter the nerve endings by the carrier causing intraterminal transfer of dopamine. The elevated external Naþ concentrations force the transporter into reverse‐mode operation and dopamine is extruded out from the nerve terminals. Dopamine released into the synaptic cleft will activate postsynaptic D1‐like (D1 and D5) and D2‐like (D2, D3, and D4) receptors. Activation of the corresponding Gs proteins by D1‐like receptors or Gi/o proteins by D2‐like receptors results in either stimulation or inhibition of adenylate cyclase and increase or decrease of cAMP production. D1 receptor stimulation acts to phosphorylate the phosphoprotein DARPP‐32 via cAMP and protein kinase A, pDARPP‐32 will then inhibit protein phosphatase‐1 (PP‐1) increasing ion channel and receptor phosphorylation. In cells expressing D2‐like receptors, Ca2þ entry activates calcineurin, which in turn, leads to dephosphorylation of pDARPP‐32

whereas dopamine in cytoplasmic store is less protected. Vesicles also retard dopamine from diffusion out of the neuron. Moreover, storage vesicles are ready for fusion with the cellular membrane and they undergo subsequent exocytosis (Hammond, 1996). Vesicular storage of dopamine serves as a depot from where dopamine can be released by appropriate physiological stimuli. The granules that store dopamine also contain ATP but in case of vesicular dopamine release, no enzyme is coreleased.

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7

Dopamine storage in and efflux from neuronal tissue pools can be characterized by the de Langer–Mulder compartmental analysis (de Langen and Mulder, 1979) by using [3H]dopamine to label (> Figure 7-3). Dopamine pools vary in their turnover rate and turnover time and also in the rate constant of release occurs . Figure 7-3 Compartmental analysis of dopamine pools in rat striatum where [3H]dopamine is taken up to and released from. The release of [3H]dopamine (kBg/g/fraction) was plotted in function of tissue [3H]dopamine content (kBq/g). Desaturation curves of [3H]dopamine efflux with different slopes yield more than on compartment in the release process. Distributional analysis revealed that the efflux compartment consists of various pools with different rate constant, turnover rate and turnover time; these stores participate in the fast, slow and resting release. Striatal slices were prepared from rat brain, loaded with [3H]dopamine and superfused. The slices were stimulated (S) electrically (20 V, 2 Hz, 2‐msec for 2 min) in fraction 4 (S). The radioactivity released from the tissue and that remained in the tissue was determined by liquid scintillation spectrometry

from them. In brain slices, accumulated [3H]dopamine distributes mainly in two compartments: an efflux compartment where release of [3H]dopamine originates from and a bound fraction, which contributes to the efflux with a limited rate (slowly exchanging compartment). Further analysis of radioactivity stored in brain tissue after incubation indicates that the efflux compartment consists of at least three different pools of [3H]dopamine from which fast and slow efflux occur in response to stimulation and a third one, which represents the source of resting dopamine outflow (Harsing, 2006) (> Figure 7-3). Other kinetic analysis has indicated that dopamine is compartmentalized into three separate pools within the presynaptic nerve terminal (Justice et al., 1988). These intraneuronal compartments are the cytosolic dopamine and two vesicular compartments. Dopamine is present in low concentrations in the cytosol where it is subject to metabolism by MAOs; their inhibitors enhance dopamine levels in this compartment. Cytosolic dopamine pool has a pivotal role in distributing newly synthesized dopamine into vesicular storage. From this store, dopamine can be released by reverse‐mode operation of dopamine transporter. There are two sources of cytosolic dopamine: one is uptake from the synaptic cleft and the other is diffusion of dopamine from the vesicular stores. Cytosolic dopamine is lost in three processes: uptake into vesicles, metabolism by MAOs to DOPAC, and efflux into the extracellular fluid. One vesicular compartment is designated as releasable bound dopamine from where action potential‐ induced depolarization evokes transmitter release. The other is a larger inactive compartment, which communicates with the active releasable compartment. The releasable compartment represents the vesicles located near the presynaptic membrane. The rapidly turning over pool contains 5–20% of the total dopamine content in the striatum. The curve of disappearance of dopamine following a‐methyl‐p‐tyrosine administration also reveals the existence of two distinct and separate phases of dopamine decline: an initial

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and rapid one and a much slower, long‐lasting one starting after drug administration (McMillen et al., 1980). These multiexponential curves of dopamine disappearance observed after a‐methyl‐p‐tyrosine indicate that dopamine is not homogeneously stored in the dopaminergic axon terminals. The biphasic decline in dopamine tissue concentrations is separated by a brief interval when dopamine levels rose to refill the rapidly turning over pool. Dopamine store with fast turnover rate preferentially releases dopamine by depolarization and the other with slow turnover rate considered as an inactive form of dopamine stored. Additional dopamine stores are those present in synaptic cleft and in extrasynaptic space, the latter can be sampled by microdialysis. The main storage site for dopamine in the dendrites may be the smooth endoplasmatic reticulum where dendritic dopamine release may occur from (Bergquist and Nissbrandt, 2005). Dopamine is also present in glial cells.

4.1 Vesicular Monoamine Transporter Accumulation of dopamine in the vesicles depends on the operation of the VMAT (Weihe and Eiden, 2000). VMAT belongs to the intracellular transporters, solute carrier (SLC)18‐gene family (Gether et al., 2006). Two VMATs were identified, one is found in the adrenal medulla and the other is present in the central nervous system, in catecholamine and serotonin neurons. The vesicular uptake process has a low substrate specificity and a variety of biogenic amines including tryptamine, tyramine, and amphetamines can be transported. These amines may compete with endogenous catecholamines for vesicular storage sites. The intravesicularly stored dopamine exists in a complex with ATP and the acidic proteins, chromogranins. The mechanism that concentrates dopamine within the vesicles is an ATP‐dependent process and it is linked to a proton pump. The driving force for uptake into synaptic vesicles is a proton electrochemical gradient generated by a vacuolar Hþ‐ATPase in the synaptic vesicle membrane. The transporter proteins have 12 transmembrane domains and are homologous to a family of bacterial drug resistance transporters. VMAT‐2 has a high affinity to reserpine, which irreversibly blocks vesicular uptake in vivo. Reserpine and related compounds (tetrabenazine, benzquinamide) also inhibit dopamine uptake into the storage vesicles and deplete available stores of dopamine. Reserpine induces depression in human due to depletion of neuronally stored catecholamines and serotonin.

5

Dopamine Plasma Membrane Transporter

Dopamine plasma membrane transporter (DAT) is a transmembrane protein, which effectively removes dopamine from synaptic cleft and returns it into the presynaptic terminals. Dopamine transporter belongs to the Naþ–Cl‐coupled transporters, SLC6‐gene family (Gether et al., 2006). The reuptake of dopamine limits its duration of action on pre‐ and postsynaptic receptors and also its diffusion to other synapses within the biophase. Moreover, the uptake process also allows the recycling and reuse of nonmetabolized dopamine molecules in the neurotransmission process. Reuptake of released dopamine by neurons is the major mode of inactivation.

5.1 Structure of Dopamine Transporter Human dopamine transporter consists of 620 amino acids. The genes for transporters responsible for uptake of dopamine have been cloned revealing protein that belongs to a larger family of neurotransporters. The protein is thought to have 12 transmembrane domains with intracellularly oriented N and C termini and a large glycosylated extracellular loop between transmembrane domains 3 and 4. Dopamine transporter possesses two to four extracellular N‐linked glycosylation sites. Domains 1, 2, and 4–8 may be involved in moving the transmitter across the membrane. The transporter is substrate for protein kinase C‐dependent phosphorylation, which reduces its activity. The dopamine transporter is phosphorylated on the N terminal tail but there are other phosphorylation sites for protein kinase A, protein kinase C, and Ca2þ/calmodulin protein kinase as well (Vaughan, 2004).

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Dopamine transporter gene expression occurs in brain areas in which dopamine is synthesized: the substantia nigra and the ventral tegmental area, it is less prevalent in the arcuate nucleus, olfactory bulb, and the retina. It is most commonly translated in cell bodies and transported to dendrites and axon fibers. Regional distribution of the carrier follows the expected localization of distinct dopamine neurons; however, dopamine transporter is not expressed in all dopamine neurons. The tuberoinfundibular dopamine cells in the hypothalamus that release dopamine into the pituitary portal blood stream, lack demonstrable dopamine transporter mRNA and protein. Dopamine transporter expression is also low in primate prefrontal cortex and a substantial amount of dopamine released is taken up by noradrenergic terminals (Gresch et al., 1995). There is dopamine reuptake into glial cells, and the functional significance of the glial reuptake remains, however, unknown.

5.2 Operation of Dopamine Transporter Dopamine transporter is saturable and its operation can be characterized by the Km and Vmax values determined from the Michaelis–Menten kinetics. The uptake process is energy‐dependent since it can be inhibited by incubation at a low temperature or by metabolic inhibitors. The neuronal reuptake is saturable and depends on Naþ cotransport as well as requiring extracellular Cl (Norregaard and Gether, 2001). Because reuptake depends on cocoupling to the Naþ gradient across the neuronal membrane, drugs such as ouabain, which inhibit Naþ–Kþ‐ATPase, inhibit the reuptake process. Veratridine, which opens Naþchannels, also inhibits the operation of the carrier. The linkage of uptake to the Naþ gradient may have physiological importance since transport is temporarily suspended at the time of depolarization‐induced release of dopamine. Coupling of transporter function to Naþ flow may lead to local changes in the Naþ gradient across the plasma membrane and thereby it can paradoxically extrude dopamine from the nerve endings. Thus, dopamine transporter may act in reverse‐mode operation, a process that conveys dopamine out of the neurons (Gainetdinov et al., 2002). The membrane transporter is not Mg2þ‐dependent, this characteristics distinguishes the neuronal membrane transporters from the vesicular transporters.

5.3 Drugs Acting on Dopamine Transporter Dopamine transporter has limited substrate specificity. Amphetamine and related drugs (methamphetamine, phenmetrazine) are taken up and force transporters actively pump dopamine out from the terminals (Norregaard and Gether, 2001). The amphetamine‐related compounds act as substrate for this transporter and thus, they compete with dopamine reuptake, and are direct releaser also. Other drugs (methylphenidate, nomifensine, amfonelic acid) block dopamine uptake but possess no dopamine releasing effects in vitro in brain slices or synaptosomal preparations. Cocaine binds to the carrier and blocks reuptake of synaptically released dopamine. The cocaine‐binding site in dopamine transporter is distinct from the substrate recognition site. Cocaine and amphetamine exert their effects on arousal by increasing extracellular dopamine concentrations. Some antidepressant and psychostimulant agents block dopamine transporter. Chronic administration of inhibitors alters the number of transporter sites. The membrane transporter is insensitive to reserpine.

6

Dopamine Receptors

Synaptically released dopamine that is not degraded enzymatically, or transported back into the presynaptic cell may activate dopamine receptors. Numerous dopamine receptors have been identified (> Table 7-2), whereas only one transporter has been cloned to dopamine. Dopamine receptors may be located on dendrites and cell bodies of neurons but also occur on axons or nerve terminals. Activation of dopamine receptors may cause decrease of dopamine release or lead to decrease or increase of various other neurotransmitters.

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. Table 7-2 Types of dopamine receptors and their agonist and antagonist ligands Receptor assays Dopamine D1 Dopamine D2

Agonist SKF38393 SKF81297 SKF82958 Bromocriptine Lisuride

Dopamine D3

BP897 Quinpirole

Dopamine D4

PD168077

Dopamine D5

SKF38393

Antagonist SCH23390

G protein Gs

Haloperidol Domperidone SKF83566 Nafadotrine U99194A Raclopide NGD941 L745870 Clozapine SCH23390

Gi/Go

Gi/Go Gi/Go Gs

Transduction coupling Stimulation of cAMP production Inhibition of cAMP production Inhibition of cAMP production Inhibition of cAMP production Stimulation of cAMP production

6.1 Classification of Dopamine Receptors The first classification of dopamine receptors into two types was proposed based upon a combination of biochemical and pharmacological criteria (Kebabian and Calne, 1979). Two types of dopamine receptors were identified based on differences in their drug specificities and signaling mechanisms. Recent evidence indicated that members of the dopamine receptor family can be generally classified as either D1‐like or D2‐ like receptors. The dopamine D1 receptor was mainly defined as the receptor associated with adenylate cyclase activation in striatal and retinal membranes and displaying low affinity for some antipsychotic drugs, such as sulpiride. The dopamine D2 receptor was defined as being associated with inhibition of prolactin release and displaying high affinity for all antipsychotic agents using radioligand binding experiments. Molecular cloning identified multiple D1 and D2‐like receptors (The IUPHAR Compendium of Receptor Characterization and Classification. 2000). According to our current view, the effects of dopamine are mediated through interaction with five different receptors usually referred to as D1 and D5 receptors and D2, D3, and D4 receptors. D1‐like receptors comprise the D1 and D5 receptors, both exhibit similar pharmacology and activate adenylate cyclase via coupling to a Gs protein and activation of protein kinase A. Subsequently, D1‐like receptors have a high affinity for benzazepines like SCH‐23390 and exhibit low affinity for benzamides (sulpiride). The D2‐like receptors compose D2, D3, and D4 receptors. Molecular cloning has demonstrated the presence of two isoforms of D2 receptors, designated D2long and D2short (Giros et al., 1989). Like D2 receptors, the receptors of type D3 also exist in different isoforms. D2‐like receptors are with similar pharmacology; they inhibit adenylate cyclase via coupling to Gi/Go proteins. D3 receptors express relatively high affinity for atypical antipsychotics and for dopamine autoreceptor inhibitors [(þ)UH‐232, (þ)AJ‐76] while D4 receptors have high affinity for clozapine. D3 receptors may in part modulate the synthesis and release of dopamine in striatum and mesolimbic regions. Some antipsychotics exhibit high to moderate affinity to D4 dopamine receptors.

6.2 Structure of Dopamine Receptors All dopamine receptor subtypes are members of the large G protein‐coupled receptor superfamily, which is characterized by seven transmembrane hydrophobic domains, an extracellular N‐ and an intracellular C terminus. Dopamine receptors contain one aspartic acid and two serine residues in transmembrane

Dopamine and the dopaminergic systems of the brain

7

domains 3 and 5, which may bind the amino and hydroxyl groups of dopamine. These receptors are subject to posttranslational modifications including glycosylation, palmytoilation, and phosphorylation. The glycosylation sites at asparagines in the N terminus are essential for the transport of receptor protein through the cell and proper folding within the plasma membrane. Cysteines in the first and second extracellular loops form disulfide bond that has importance to maintain the three‐dimensional structure of the protein within the membrane. Sequences of phosphorylation are found in the second and third intracellular loops and the C terminal tail of the receptor. D1 receptors possess small i3 loops and long C‐tails and D2 receptors have large i3 loops and short C‐tails. Dopamine receptors form a packet‐like structure in which dopamine is recognized and bound. The binding of dopamine to a membrane‐bound receptor initiates a conformational change in the receptor such that it alters a G protein, which in turn is coupled to ion channels or second messenger systems. Two regions in the third intracellular loop are essential for binding of receptors to G protein a subunit.

6.3 Postsynaptic Dopamine Receptors Response to postsynaptic D1 and D2 receptor stimulation either activates or inhibits a messenger cascade involving the phosphorylation of a dopamine‐ and cAMP‐regulated phosphoprotein, DARPP‐32 (Greengard et al., 1999). The DARPP‐32 signaling pathway has a central role in mediating signal transduction within neurons like GABAergic medium‐sized spiny neurons in the striatum. Stimulation of D1 receptors by dopamine acts via cAMP and protein kinase A to phosphorylate phosphoprotein DARPP‐32, which in turn inhibits the activity of protein phosphatase‐1 (PP‐1). Besides changes in signal transduction system, electrophysiological changes after D1 receptor stimulation have also been reported. Thus, D1 receptor stimulation in the striatum reduces fast sodium conductance and N‐ and P‐type calcium currents and also enhances L‐type Ca2þ currents via a protein kinase A‐mediated process (Grace, 2002). In contrast, stimulation of D2 receptors leads to calcium stimulation of protein phosphatase‐2B. D2 receptor activation in enkephalin‐containing striatal GABAergic neurons (striatopallidal pathway) causes dephosphorylation of DARPP‐32 by Ca2þ influx‐activated calcineurin. Thus, D1 and D2 receptors exert opposite effects on centrally positioned DARPP‐32 in the signal transduction of striatal GABA neurons. Furthermore, dopamine agonists can exert excitation on GABA release within the striatum via D1 receptors and an inhibitory influence via D2 receptors as shown in a functional assay (Harsing and Zigmond, 1997).

6.4 Presynaptic Dopamine Receptors Whereas postsynaptic D2 receptors are associated with intracellular signaling, presynaptic dopamine D2 receptors regulate the release and synthesis of dopamine as well as the firing activity of dopaminergic neurons (> Figure 7-2). This regulation is primarily inhibitory in nature as activation of dopamine autoreceptors on the same presynaptic terminals can curtail the release and synthesis of dopamine, whereas those reside on cell bodies reduce neuronal firing activity. All three types of dopamine autoreceptors belong to the D2 family of dopamine receptors, which includes D2, D3, and D4 receptors. Presynaptic D2 dopamine receptors serve as autoreceptors because their activation can inhibit the cells by responding dopamine released from the same neurons; this kind of regulation is often designated as negative feedback regulation. Autoreceptors are distributed in all parts of the neurons, including the soma, the dendrites, and the nerve terminals. The release‐mediating dopamine autoreceptors at the nerve terminal respond to the transmitter released into the synaptic cleft, and those located on the cell body may respond to dendritic dopamine release. Release‐mediated autoreceptors are coupled to Gi protein and they dampen further release of dopamine by one or more of the following mechanisms (Bagdy and Harsing, 1995; Nestler et al., 2001): 1. Inhibitions of G protein‐coupled voltage‐sensitive Ca2þ channels that leads to restriction of available Ca2þ for depolarization.

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Dopamine and the dopaminergic systems of the brain

Activation of the inwardly rectifying Kþ channels, which may increase potassium ion permeability through the cell membrane. Inhibition of presynaptically located receptor‐coupled adenylate cyclase, which is mediated by Gi/o protein. This may cause inhibition of protein kinase A that regulates proteins involved in neurotransmitter release.

D2 autoreceptors with different functions may be coupled to distinct transduction mechanisms. The released dopamine acts homeostatically at the synthesis‐modulating autoreceptors to control neurotransmitter synthesis. Dopamine receptors directly regulate dopamine synthesis: dopamine agonists decrease and antagonists increase synthesis of the neurotransmitter. Synthesis‐modulating autoreceptors are not present on all neurons; some midbrain dopamine neurons projecting to the prefrontal cortex appear to lack synthesis‐modulating autoreceptors. Synthesis‐regulating dopamine autoreceptors may belong to D3 rather than the D2 subtype in the striatum (Meller et al., 1993). Dopamine‐containing neurons in the midbrain exhibit spontaneous firing that is driven by an endogenous pacemaker conductance with their activity modulated by autoreceptors and afferent inputs. One of the prominent regulators of dopamine neuronal activity is the impulse‐mediating dopamine autoreceptor. These autoreceptors are located on the soma and dendrites of dopamine neurons. Impulse‐mediating autoreceptors are believed to exert a tonic downregulation of dopamine neuron activity, maintaining their firing rate within a stable range of activity. Somatodendritic autoreceptors are stimulated by an extracellular pool of dopamine released from the dendrites of the same or neighboring dopamine neurons. Dopamine agonists inhibit spike firing and corresponding antagonists reverse this effect.

6.5 Regional Distribution of Dopamine Autoreceptors Assessed by the g‐Butyrolactone Model g‐Butyrolactone (GBL) increases the concentrations of dopamine in several regions of the rat brain (Roth, 1984). This increase in dopamine levels is due to inhibition of impulse flow in dopamine neurons and the reduced release leads to disinhibition of synthesis‐mediated autoreceptors. The GBL‐evoked increase in dopamine synthesis can be further enhanced with inhibition of dopa decarboxylase elicited by concomitant administration of NSD‐1015 (Harsing and Vizi, 1991). The increase in dopamine levels after GBL injection can be antagonized by the D2 receptor agonist apomorphine in those brain areas where dopaminergic terminals possess dopamine‐sensitive autoreceptors. The use of this technique led to differentiate among dopamine nerve terminals: as those in the striatum, olfactorial bulb, amygdala, and piriform cortex express dopamine‐sensitive autoreceptors, whereas they are absent in dopaminergic nerve terminals of the prefrontal, cingulated, and entorthinal cortices. GBL pretreatment also decreases the formation of 3‐methoxytyramine, an indirect index for dopamine release in rat striatum (Westerink and Spaan, 1982).

6.6 Changes in Dopamine Receptor Sensitivity and Expression Receptor desensitization may occur when transmitter no longer causes a cellular response (Kuhar et al., 2006). Chronic agonist stimulation may induce this phenomenon. Desensitization may be due to receptor phosphorylation by protein kinase A, protein kinase C, or G protein‐coupled receptor kinases. A related phenomenon is termed downregulation, which usually occurs with a slower time course and involves cellular adaptations such as receptor degradation. MAO inhibitors and tricyclic antidepressants that increase concentrations of dopamine within the synaptic cleft leads to functional subsensitivity. Partial lesion of the dopamine system results in dopamine receptor supersensitivity in the remaining neurons. Destruction of neurons with neurotoxins such as 6‐hydroxydopamine evokes functional supersensitivity. Supersensitivity can result from a rapid loss of release sites following denervation. Chronic

Dopamine and the dopaminergic systems of the brain

7

administration of antagonists increases the density of D1 and D2 receptors, this increase in postsynaptic sites occurs in a longer time course. Supersensitivity of dopamine receptors that have been chronically blocked during treatment of patients with first generation antipsychotic drugs leads to development of excessive motor activity called tardive dyskinesia.

7

Dopamine Release from Neuronal Stores

Dopamine release may occur from axon terminals or dendrites of neurons by action potential propagation or reverse‐mode operation of the plasma dopamine transporter. Agonists of ion channel‐coupled receptors (glutamate or nicotinic receptors) may also increase neurotransmitter outflow from dopamine neurons. The mechanisms of these processes are different.

7.1 Action Potential Propagation‐Induced Dopamine Release Membrane depolarization induced by action potential propagation evokes dopamine release from synaptic vesicles located in nerve endings (> Figure 7-4). This release occurs from storage vesicles by the

. Figure 7-4 The time course of [3H]dopamine release measured from striatal slices. Slices were prepared from rat brain, loaded with [3H]dopamine and superfused with Krebs‐bicarbonate buffer. The slices were stimulated electrically (20 V, 2 Hz, 2‐msec for 2 min in fractions 4(S1) and 15(S2)) and the fractional release of [3H]dopamine (i.e., a percentage of the amount of [3H]dopamine in the tissue at the time of the release) was calculated. The calculated ratio of fractional release S2 over fractional release S1 (S2/S1) was 1.021. The radioactivity released from the tissue and that remained in the tissue was determined by liquid scintillation spectrometry

mechanism of exocytosis. Vesicular dopamine release is an external Ca2þ‐dependent process, as increase in free Ca2þ concentrations in the cytosol triggers fusion of secretory vesicles with plasma membrane and lack of free Ca2þ abolishes it. The rise of intracellular free Ca2þ is a consequence of Ca2þ entry through plasma membrane Ca2þ channels and other entry processes like opening of receptor‐coupled ion channels

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permeable to bivalent cations. Release of Ca2þ from intracellular stores may have less importance in the exocytotic dopamine release. Membrane depolarization induced by elevated external potassium concentration leads to increase in dopamine release and this release also needs the presence of external Ca2þ. Dopamine release evoked by electrical stimulation, which mimics action potential propagation, can be abolished by tetrodotoxin, a drug which blocks voltage‐dependent sodium channels. On the contrary, tetrodotoxin does not abolish dopamine release evoked by high potassium depolarization, a stimulation procedure confines to the plasma membrane of the nerve terminals. Dopamine release that occurs typically by a Ca2þ‐dependent process is also designated as phasic release (Grace, 1991). The compartment, where phasic dopamine release originates from, can be characterized by carbon fiber voltammetry that allows to measure dopamine release in a real time base. Dopamine released from nerve ending with a phasic manner is removed from the synaptic cleft by the process of reuptake. The reuptake process does not capture all dopamine released as some may diffuse out from the synaptic cleft establishing dopamine pool in the biophase. Dopamine present in the extrasynaptic space represents the bases of nonsynaptic dopaminergic neurotransmission (Vizi, 2000) and it is often called tonic dopamine. Extrasynaptic compartments of tonic dopamine can be sampled by brain microdialysis, a technique that exhibits slower measures of dopamine dynamics. Phasic versus tonic dopamine release has been shown to have importance in normal and dysfunctional dopamine regulations related to certain psychiatric disorders such as schizophrenia, attention deficit hyperactivity disorder or drug abuse (Grace, 1991). From dendrites, dopamine can be released through a process that may not necessarily involve conventional exocytosis, i.e., the release, according to some observations, is not Ca2þ‐dependent.

7.2 Dopamine Release Evoked by Reverse‐Mode Operation of Dopamine Transporters Dopamine active transport could be bidirectional and able to evoke release by the same exchange‐diffusion process as that involved in the uptake function. Amphetamine and its analogs, methamphetamine and phenmetrazine, evoke direct release of dopamine and also inhibit its reuptake; these effects can be demonstrated both in vivo and in vitro. The releasing effect of amphetamine and its derivatives is mediated by outward transport of dopamine (Leviel, 2001). The releasing molecules that evoke dopamine release by reverse‐mode operation of the transporter induce the following steps: 1. Dopamine releaser drugs enter axon terminals by an operation of the transporter in the normal mode. 2. The releaser molecules evoke intraterminal transfer of dopamine from an electrically releasable pool to cytosolic compartment and thus dopamine concentrations increase within the terminals. 3. Dopamine is then transported out from the nerve terminals by reverse‐mode operation of the transporter. The release of dopamine by exchange‐diffusion is temperature‐ and Naþ‐dependent, saturable, and stereoselective. Exocytosis requires extracellular Ca2þ but dopamine transporter‐mediated outward transport does not. This effect is also not receptor‐mediated. Protein kinase C is involved in the external Ca2þ‐ independent dopamine release: activators of protein kinase C result in increase of dopamine efflux even in the absence of external Ca2þ (Gnegy, 2003). (þ)Amphetamine releases dopamine from a pool that is insensitive to reserpine but dependent on newly synthesized dopamine (i.e., a‐methyl‐p‐tyrosine sensitive). Other classes of drugs that inhibit dopamine reuptake (methylphenidate, cocaine, and amphenolic acid) do not induce dopamine release in vitro but they block the dopamine releasing effect of (þ)amphetamine. Methylphenidate evokes ‘‘neurogenic overflow’’ of dopamine in vivo and this effect may be explained by increased exchange of dopamine from the large, reserpine‐sensitive storage pool, too much smaller releasable sites (McMillen, 1983). This effect of methylphenidate has first been demonstrated as a potentiation of dopaminergic drugs in animal behavioral tests.

Dopamine and the dopaminergic systems of the brain

7

7.3 Dopamine Release Evoked by Ion Channel‐Coupled Receptors Ligands for ion channel‐coupled receptors (glutamate, nicotinic) can release dopamine by a mechanism that involves reverse‐mode operation of dopamine transporter and, in some cases, vesicular process as well (Lonart and Zigmond, 1991). Thus, glutamate is taken up into dopamine nerve endings by heterotransporter and Naþ is cotransported during this process. This leads a rise in intracellular Naþ concentrations, which promotes reduced membrane potential across nerve terminals. Enhanced intracellular Naþ concentrations shift dopamine transporter operation into reverse mode resulting in an increase in dopamine efflux from the nerve terminals. In addition, activation of ionotropic glutamate receptors permits influx of Naþ through receptor‐coupled ion channels and the rise of intracellular Naþ causes further reversal of Naþ‐dependent dopamine transport with a consequent efflux of dopamine from cytoplasmic stores. Moreover, excitatory amino acids also induce Ca2þ entry through receptor‐coupled ion channels followed by a further increase in intracellular Ca2þ concentrations due to opening of voltage‐dependent Ca2þ channels. The rise in intracellular Ca2þ concentrations may then be utilized for dopamine release process originating from nerve ending neurotransmitter stores. Activation of nicotinic receptors by agonists may also directly open ligand‐gated ion channels that are permeable to monovalent cations and also Ca2þ (Wonnacott et al., 1995). When nicotine binds to and activates nicotinic receptors, Naþ enters the cells through nicotinic receptor‐coupled ion channels inducing local membrane depolarization. Activation of nicotinic receptors may elevate free Ca2þ intracellularly in an indirect way by depolarizing the cell membrane enough for Ca2þ entry through voltage sensitive Ca2þ channels. As a consequence of the enhanced free intracellular Ca2þ, an exocytotic process may trigger dopamine release from vesicular pool. In many experimental conditions, dopamine release evoked by nicotine has been reported to be partly external Ca2þ‐dependent (Harsing et al., 1992).

7.4 Regulation of Dopamine Release The activity of dopaminergic neurons is determined by at least three different events. These are (1) the spontaneous discharge activity of dopamine neurons, (2) the autoinhibitory properties that include regulation of release, synthesis, and neural firing rate, and (3) excitatory and inhibitory afferent inputs mediated by heteroreceptor population of dopamine neurons. The glutamate‐ and GABA heteroreceptor‐ mediated regulation of dopamine release is particularly well characterized in the striatum. In addition, dopamine release can also be controlled by number local factors such as nitric oxide (NO) that is located released from striatal interneurons. When added, substrates for nitric oxide synthase (NOS) or NO generator compounds stimulate dopamine release in a Ca2þ‐dependent fashion (West and Galloway, 1997). The NO system and excitatory amino acids may interact with dopamine neuronal firing to regulate dopamine release from presynaptic sites in the striatum.

8

Dopaminergic Innervations in the Central Nervous System

A great portion of dopaminergic neurons in the central nervous system is located in three dopaminergic nuclei, the substantia nigra pars compacta (area A9), the ventral tegmental area (area A10), and the arcuate nucleus. Pathway originating from the substantia nigra pars compacta projects to the caudate nucleus and putamen and it forms the nigrostriatal dopaminergic system. Neurons from the ventral tegmental area largely project to the limbic structures like nucleus accumbens, prefrontal cortex, and cingulate cortex. These projections are designated as mesocortical/mesolimbic dopaminergic pathways. From the hypothalamic arcuate nucleus, dopaminergic neurons project to the pituitary gland, this system is designated as the tuberoinfundibular dopaminergic system. The tuberoinfundibular intermediate‐length dopaminergic system controls prolactin release from the anterior pituitary and its blockade by antipsychotics leads to

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neuroendocrine effects (hyperprolactinemia or lactation) characteristic for this class of drugs. Other dopaminergic neurons distribute in the retina (ultrashort dopaminergic system) and other areas like the olfactory bulb and the lemniscal area (Cooper et al., 1996).

8.1 Dopamine in the Striatum In the neostriatum, D1 receptors are predominantly present in the GABAergic striatonigral GABAergic neurons, which contain substance P and dynorphin as cotransmitters, and project to the substantia nigra. D2 receptors occur mainly in striatopallidal GABAergic neurons, these neurons also contain enkephalins. Dopamine exerts inhibitory action on corticostriatal glutamatergic axon terminals, and this effect is mediated by D2 receptors (Harsing and Vizi, 1991). Striatal dopamine release controls movement patterns in physiological conditions and voluntary movement is impaired when dopaminergic tone is impaired by dopamine cell loss. Dopamine acting on D1 and D2 receptors influences two opposite types of synaptic plasticity, in the striatum, the long‐term potentiation and long‐term depression. Such plasticity within the striatum may be involved in acquisition of complete motor skills. The nigrostriatal neurons are the neural substrate for antiparkinsonian drugs and antipsychotic agents induce extrapyramidal side effects.

8.2 Dopamine in the Cerebral Cortex Cortical pyramidal neurons in layers V and VI receive dopaminergic influence from the ventral tegmental area and the released dopamine acts on D1/D5 receptors on apical dendrites of the pyramidal neurons. Dopamine terminals in the prefrontal cortex do not contain dopamine transporters (Lewis and Gonzales‐ Burgos, 2006). As a consequence, dopamine released from these sites would be free to diffuse to a much greater extent. The mesocortical and mesolimbic dopaminergic neurons are involved in cognitive and emotive functions and in the pathophysiology of various forms of psychosis. These dopaminergic systems are widely used to explain the mode of action of antipsychotic drugs and chemicals inducing drug abuse. The reward pathway is also well characterized: dopaminergic connection between the ventral tegmental areas and the nucleus accumbens mediating reinforcing properties of drugs of abuse. Stress causes dopamine release in the amygdala and lesion of amygdala tends to block stress‐induced increases in dopamine levels in the prefrontal cortex.

9

Altered Dopaminergic Neurotransmission

9.1 Neurotoxins Used for Destruction of Dopaminergic Neurons The dopamine analog 6‐hydroxydopamine, when it is administered directly into brain tissue, is selectively transported into dopaminergic axon terminals via high affinity uptake system. 6‐hydroxydopamine is then readily oxidizes to form of a series of cytotoxic compounds, 6‐hydroxydopamine‐p‐quinone, hydrogen  peroxide (H2O2), superoxide anion (O 2 ), and hydroxyl radical (OH ) (Zigmond et al., 1992). The accumulation of potentially cytotoxic compounds destroys dopaminergic elements by lipid peroxidation and protein and DNA damage. Dopamine uptake inhibitors can suspended the neurotoxic effect of 6‐hydroxydopamine whereas the MAO inhibitor pargyline potentiates it. Neurochemical changes after 6‐hydroxydopamine‐induced lesion involve increase dopamine release, synthesis and turnover rate, and reduced dopamine uptake (Zigmond, 1990; Juranyi et al., 2004). The 6‐hydroxydopamine‐induced destruction of dopamine neurons leads to severe neurological deficits, such as akinesia, reduce food and water intake, and lack of response to sensory stimuli. Dopamine agonists or muscarinic antagonists can reverse akinesia.

Dopamine and the dopaminergic systems of the brain

7

1‐Methyl‐4‐phenyl‐1,2,3,6‐tetrahydropyridine (MPTP) is a selective neurotoxin for dopaminergic neurons, which is commonly used to induce experimental Parkinson’s disease in mice and monkeys (Heikkila and Sonsalla, 1987). MPTP enters the brain and B form of MAO converts it to 1‐methyl‐4‐ phenylpyridinium (MPPþ) (Heikkila et al., 1984). This product is then taken up into dopaminergic nerve terminals by dopamine transporters and it either inhibits complex II of the respiratory chain or generate oxygen‐reactive species within the nerve endings (Marsden, 2006). MPTP‐induced degeneration of dopaminergic neurons can be suspended with inhibition of MAO‐B by L‐deprenyl or by selective dopamine uptake blockers, mazidol or nomifensine (Javitch et al., 1985). Knockout mice lacking dopamine transporter express no sensitivity to the neurotoxin MPTP.

9.2 Dopaminergic Neurotransmission in Knockout Mice Tyrosine hydroxylase knockout mouse is not viable, whereas mice lacking dopamine decarboxylase exhibit hypersensitive to amphetamine. It was reported that MAO and catecol‐O‐methyl‐transferase knockout mice show aggression and altered anxiety behavior. D2 receptor‐deficient animals showed reduced dopamine autoreceptor‐mediated cell firing inhibition. Absence of dopamine transporter in mice induces hyperactivity, which may be the consequence of hyperdopaminergic state (Gainetdinov et al., 2002). In dopamine transporter knockout mice, there is an increase in extracellular dopamine levels and delayed clearance of dopamine released, and increased dopamine synthesis was observed. Because there is a loss of autoreceptor‐ mediated tone, tyrosine hydroxylase is disinhibited due to a lack of intraneuronal dopamine and dopamine turnover is markedly increased. These changes in dopaminergic neurotransmission in knockout mice are similar to the normal function of prefrontal dopaminergic neurons.

10 Conclusions and Future Avenues The success in dopamine research is due to the fact that the pathology of a series of psychiatric and neurological disorders can be explained based upon their dopamine theory. The positive symptoms of schizophrenia (hallucination, delusion, thought disorder) and cognitive deficits characteristics for this disorder can satisfyingly be explained by hyperfunctionality of subcortical dopaminergic systems and a deficit in cortical dopaminergic neurotransmission (Lewis and Gonzales‐Burgos, 2006). Both the first and the second generations of antipsychotic drugs exert potent antagonistic effects on D2 dopamine receptors although the ratio of 5‐HT2A versus D2 receptor antagonism is more pronounced for the second generation antischizophrenic agents. Currently used antipsychotic agents exhibit a wild range of side effects due to the broad range of receptors on which these agents act. Drugs acting preferentially at D3 binding site (S33138, Millan et al., 2002) or as D2/D3 receptor antagonists/partial agonists (RGH‐188, Kiss et al., 2006) or D2/D3 antagonist with D4 partial agonistic effect (F15063, Newman‐Tancredi et al., 2006) represent a series of third generation antipsychotic compounds. Some of these drugs are now in Phase I/Phase II human clinical trials. It is strongly believed that compounds with these receptor‐binding profiles will lead to antipsychotic activity associated with lower incident of adverse side effects. Of the neurodegenerative disorders, Parkinson’s disease is characterized with loss of dopaminergic neurons in the substantia nigra pars compacta. The reduced dopaminergic tone and the consequent disbalance between the direct and indirect GABAergic projection neurons in the striatum may satisfyingly explain the symptoms of Parkinson’s disease (tremor at rest, bradykinesia, and muscle rigidity). The impaired dopaminergic influence may be enhanced by addition of the dopamine precursor L‐dopa in combination with the peripheral dopa decarboxylase inhibitor carbidopa. Besides supplementary therapy, inhibition of dopamine breakdown can also be of therapeutic value: the B type MAO inhibitor L‐deprenyl is used in the therapy of Parkinson’s disease for this purpose. Rasagiline, another B type MAO inhibitor, has also been shown to be effective in treatment of Parkinson’s disease (Siderowf, 2002).

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Acknowledgement This work was supported in part by grants of the Hungarian Research Foundation (OTKA T‐43511) and Hungarian Medical Research Council (ETT‐482/2003). The author acknowledges the editorial work of Ms. Judit Puskas.

References Bagdy E, Harsing LG Jr. 1995. The role of various calcium and potassium channels in the regulation of somatodendritic serotonin release. Neurochem Res 20: 1409-1415. Bergquist F, Nissbrandt H. 2005. Dopamine release in substantia nigra: Release mechanisms and physiological function in motor control. Dendritic Neurotransmitter Release. Ludwig M, editor. New York: Springer; pp. 85-99. Cooper JR, Bloom FE, Roth RH. 1996. Dopamine. The Biochemical Basis of Neuropharmacology. Oxford University Press; New York: pp. 293-351. Dahlstrom A, Fuxe K. 1964. Evidence for the existence of monoamine containing neurons in the central nervous system. 1. Demonstration of monoamines in the cell bodies of brain stem neurons. Acta Physiol Scand Suppl 232: 1-55. de Langen CDJ, Mulder AH. 1979. Compartmental analysis of the accumulation of 3H‐dopamine in synaptosomes from rat corpus striatum. Naunyn Schmiedebergs Arch Pharmacol 308: 31-39. Demarest KT, Smith DJ, Azzaro AJ. 1980. The presence of the type A form of monoamine oxidase within nigrostriatal dopamine‐containing neurons. J Pharmacol Exp Ther 215: 461-468. Deutch AY, Roth RH. 2004. Pharmacology and biochemistry of synaptic transmission: Classic transmitters. From Molecules to Network. Byrne JH, Roberts JL, editors. New York: Elsevier Academic Press; pp. 245-278. Gainetdinov RR, Sotnikova TD, Caron MG. 2002. Monoamine transporter pharmacology and mutant mice. Trends Pharmacol Sci 23: 367-373. Gether U, Andersen PH, Larsson OM, Schousboe A. 2006. Neurotransmitter transporters: Molecular function of important drug targets. Trends Pharmacol Sci 27: 375-383. Giros B, Sokoloff P, Martres MP, Riou JF, Emorine LJ, et al. 1989. Alternative splicing directs the expression of two D2 dopamine receptor isoforms. Nature 342: 923-926. Glover V, Sandler M, Owen F, Riley GJ. 1977. Dopamine is a monoamine oxidase B substrate in man. Nature 265: 80-81. Gnegy ME. 2003. The effect of phosphorylation on amphetamine‐mediated outward transport. Eur J Pharmacol 479: 83-91. Grace AA. 1991. Phasic versus tonic dopamine release and the modulation of dopamine system responsivity: A hypothesis for the etiology of schizophrenia. Neuroscience 41: 1-24.

Grace AA. 2002. Dopamine. Neuropsychopharmacology: The Fifth Generation of Progress. Davis KL, Charney D, Coyle JT, Nemeroff C, editors. New York: Lippincott Williams and Wilkins; pp. 119-132. Greengard P, Allen PB, Nairn AC. 1999. Beyond the dopamine receptor: The DARPP‐32/protein phosphatase‐1 cascade. Neuron 23: 435-447. Gresch PJ, Sved AF, Zigmond MJ, Finlay JM. 1995. Local influence of endogenous norepinephrine on extracellular dopamine in rat medial prefrontal cortex. J Neurochem 65: 111-116. Hammond C. 1996. Neurotransmitter release. Cellular and Molecular Neurobiology. Hammond C, editor. New York: Academic Press; pp. 188-214. Harsing LG Jr, Vizi ES. 1991. Alpha 2‐adrenoceptors are not involved in the regulation of striatal glutamate release: Comparison to dopaminergic inhibition. J Neurosci Res 28: 376-381. Harsing LG Jr, Sershen H, Vizi SE, Lajtha A. 1992. N‐type calcium channels are involved in the dopamine releasing effect of nicotine. Neurochem Res 17: 729-734. Harsing LG Jr, Zigmond MJ. 1997. Influence of dopamine on GABA release in striatum: Evidence for D1‐D2 interactions and non‐synaptic influences. Neuroscience 77: 419-429. Harsing LG Jr. 2006. The pharmacology of the neurochemical transmission in the midbrain raphe nuclei of the rat. Curr Neuropharmacol 4: 313-339. Heal DJ, Frankland AT, Buckett WR. 1990. A new and highly sensitive method for measuring 3‐methoxytyramine using HPLC with electrochemical detection. Studies with drugs which alter dopamine metabolism in the brain. Neuropharmacology 29: 1141-1150. Heikkila RE, Manzino L, Cabbat FS, Duvoisin RC. 1984. Protection against the dopaminergic neurotoxicity of 1‐methyl‐4‐phenyl‐1,2,5,6‐tetrahydropyridine by monoamine oxidase inhibitors. Nature 311: 467-469. Heikkila RE, Sonsalla PK. 1987. The use of the MPTP‐treated mouse as an animal model of parkinsonism. Can J Neurol Sci 14: 436-440. Javitch JA, D’Amato RJ, Strittmatter SM, Snyder SH. 1985. Parkinsonism‐inducing neurotoxin, N‐methyl‐4‐ phenyl‐1,2,3,6‐tetrahydropyridine: Uptake of the metabolite N‐methyl‐4‐phenylpyridine by dopamine neurons

Dopamine and the dopaminergic systems of the brain explains selective toxicity. Proc Natl Acad Sci USA 82: 2173-2177. Juranyi Z, Sziray N, Marko B, Levay G, Harsing LG Jr. 2004. AMPA receptor blockade potentiates the stimulatory effect of L‐DOPA on dopamine release in dopamine‐deficient corticostriatal slice preparation. Crit Rev Neurobiol 16: 129-139. Justice JB Jr, Nicolaysen LC, Michael AC. 1988. Modeling the dopaminergic nerve terminal. J Neurosci Methods 22: 239-252. Kebabian JW, Calne DB. 1979. Multiple receptors for dopamine. Nature 277: 93-96. Kiss B, Schmidt E, Horvath A, Orosz S, Laszlovszky I, et al. 2006. Neurochemical profile of RGH‐188, a novel dopamine D3/D2 antagonist/partial agonist. Soc Neurosci 36: Abstr 93.2. Kuhar MJ, Minneman K, Muly EC. 2006. Catecholamines. Basic Neurochemistry, Molecular, Cellular, and Medical Aspects. Siegel GJ, Albers RW, Brady ST, Price DL, editors. New York: Elsevier Academic Press; pp. 211-223. Leviel V. 2001. The reverse transport of DA, what physiological significance? Neurochem Int 38: 83-106. Lewis DA, Gonzalez‐Burgos G. 2006. Pathophysiologically based treatment interventions in schizophrenia. Nat Med 12: 1016-1022. Lonart G, Zigmond MJ. 1991. High glutamate concentrations evoke Ca(þþ)‐independent dopamine release from striatal slices: A possible role of reverse dopamine transport. J Pharmacol Exp Ther 256: 1132-1138. Marsden CA. 2006. Dopamine: The rewarding years. Br J Pharmacol 147 (Suppl 1): S136-S144. McMillen BA, German DC, Shore PA. 1980. Functional and pharmacological significance of brain dopamine and norepinephrine storage pools. Biochem Pharmacol 29: 3045-3050. McMillen BA. 1983. CNS stimulants: Two distinct mechanisms of action for amphetamine‐like drugs. Trends Pharmacol Sci 4: 429-432. Megyeri K, Marko B, Sziray N, Gacsalyi I, Juranyi Z, et al. 2007. Effects of 2,3‐benzodiazepine AMPA antagonists on dopamine turnover in the striatum of rats with experimental parkinsonism. Brain Res Bull 71: 501-507. Meller E, Bohmaker K, Goldstein M, Basham DA. 1993. Evidence that striatal synthesis‐inhibiting autoreceptors are dopamine D3 receptors. Eur J Pharmacol 249: R5-R6. Millan MJ, Cussac D, Newman‐Tancredi A, Gobert A, Lejeune F, et al. 2002. The novel benzopyrano‐pyrrole and antipsychotic, S33138, is a preferential antagonist at dopamine (DA) D3 vs D2 receptors. Am Soc Neurosci 32, Abstr 894.14. Nestler EJ, Hyman SE, Malenka RC. 2001. Molecular Neuropharmacology. A Foundation for Clinical Neuroscience. McGraw‐Hill; New York: pp. 115-137.

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Newman‐Tancredi A, Assie M, Martel J, Cosi C, Heusler P, et al. 2006. F15063, an antipsychotic with D2/D3 antagonist, 5‐HT1A agonist, and D4 partial agonist properties: Activity in vitro and neurochemical profile in rodents. Soc Neurosci, 36: Abstr 93.6. Norregaard L, Gether U. 2001. The monoamine neurotransmitter transporters: Structure, conformational changes and molecular gating. Curr Opin Drug Discov Devel 4: 591-601. Ogawa N, Tanaka K, Asanuma M. 2000. Bromocriptine markedly suppresses levodopa‐induced abnormal increase of dopamine turnover in the parkinsonian striatum. Neurochem Res 25: 755-758. Roth RH. 1984. CNS dopamine autoreceptors: Distribution, pharmacology, and function. Ann N Y Acad Sci 430: 27-53. Siderowf A. 2002. A controlled trial of rasagiline in early Parkinson disease: The TEMPO study. Arch Neurol 59: 1937-1943. Snyder AM, Keller RW, Zigmond MJ. 1990. Dopamin efflux from striatal slices after intracerebral 6‐hydroxydopamine: Evidence for compensatory hyperactivity of residual terminals. J Pharm Exp Ther 253: 867-876. The IUPHAR Compendium of Receptor Characterization and Classification. 2000. 2nd edition, London: IUPHAR Media; pp. 170–181. Thony B, Auerbach G, Blau N. 2000. Tetrahydrobiopterin biosynthesis, regeneration and functions. Biochem J 347 Pt 1: 1-16. Vaughan RA. 2004. Phosphorylation and regulation of psychostimulant‐sensitive neurotransmitter transporters. J Pharmacol Exp Ther 310: 1-7. Vizi ES, Harsing LG Jr, Gaal J, Kapocsi J, Bernath S, et al. 1986. CH‐38083, a selective, potent antagonist of alpha‐2 adrenoceptors. J Pharmacol Exp Ther 238: 701-706. Vizi ES. 2000. Role of high‐affinity receptors and membrane transporters in nonsynaptic communication and drug action in the CNS. Pharmacol Rev 55: 8775-8779. Von Bohlen und Halbach O, Dermietzel R. 2002. Neurotransmitters and Neuromodulators. Wiley‐VCH Verlag GmbH; Weinheim: pp. 53-63. Walter DS, Flockhart IR, Haynes MJ, Howlett DR, Lane AC, et al. 1984. Effects of idazoxan on catecholamine systems in rat brain. Biochem Pharmacol 33: 2553-2557. Weihe E, Eiden LE. 2000. Chemical neuroanatomy of the vesicular amine transporters. FASEB J 14: 2435-2449. West AR, Galloway MP. 1997. Endogenous nitric oxide facilitates striatal dopamine and glutamate efflux in vivo: Role of ionotropic glutamate receptor‐dependent mechanisms. Neuropharmacology 36: 1571-1581. Westerink BH, Spaan SJ. 1982. On the significance of endogenous 3‐methoxytyramine for the effects of centrally acting

169

170

7

Dopamine and the dopaminergic systems of the brain

drugs on dopamine release in the rat brain. J Neurochem 38: 680-686. Wonnacott S, Gothert M, Chahl LA, Willow M, Nicholson GM. 1995. Modulation of neurotransmitter release by some therapeutic and socially used drugs. Neurotransmitter release and its Modulation. Powis DA, Bunn SJ, editors. Cambridge University Press; pp. 293–328.

Zigmond MJ. 1990. Compensations after lesions of central dopaminergic neurons: Some clinical and basic implications. Trends Neurosci 13: 290-296. Zigmond MJ, Hastings TG, Abercrombie ED. 1992. Neurochemical responses to 6‐hydroxydopamine and L‐dopa therapy: Implications for Parkinson’s disease. Ann N Y Acad Sci 648: 71-86.

8

5‐Hydroxytryptamine in the Central Nervous System

A. C. Dutton . N. M. Barnes

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 172

2

Neuroanatomy of the 5‐Hydroxytryptaminergic System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 172

3

Forebrain Projections of the Raphe Nuclei . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 172

4

The Physiology of 5‐HT Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 172

5 5.1 5.1.1 5.1.2 5.1.3 5.1.4 5.1.5 5.2 5.2.1 5.2.2 5.2.3 5.3 5.4 5.5 5.6 5.7

The 5‐HT Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 174 The 5‐HT1 Receptor Family . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 174 The 5‐HT1A Receptor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 175 The 5‐HT1B Receptor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 178 The 5‐HT1D Receptor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 181 The 5‐ht1E Receptor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 182 The 5‐HT1F Receptor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 182 The 5‐HT2 Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 183 5‐HT2A Receptor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 183 The 5‐HT2B Receptor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 185 The 5‐HT2C Receptor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 186 The 5‐HT3 Receptor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 187 The 5‐HT4 Receptor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 189 The 5‐ht5 Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 190 The 5‐HT6 Receptor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 192 The 5‐HT7 Receptor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 194

6

The 5‐HT Transporter (SERT) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 196

7

Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 198

#

2008 Springer ScienceþBusiness Media, LLC.

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5‐Hydroxytryptamine in the central nervous system

Abstract: 5-Hydroxytryptamine (5-HT, serotonin), mediates numerous physiological processes in the CNS. The diversity of function is at least partly a consequence of the 14 distinct receptors evident in mammals. The present review describes the biochemistry, physiology and pharmacology of the 5-HT system in the brain. List of Abbreviations: AHP, afterhyperpolarization potential; APP, amyloid precursor protein; BDNF, brain‐derived neurotrophic factor; EPSCs, excitatory postsynaptic currents; GPCR, G‐protein‐coupled receptor; IPSCs, inhibitory postsynaptic currents; NREMS, Nonrapid eye movement sleep; OCD, obsessive compulsive disorder; PET, positron emission tomography; PLC, phospholipase C; PPI, prepulse inhibition; REM, rapid eye movement; SAR, structure–affinity relationship; SCN, suprachiasmatic nucleus

1

Introduction

5‐Hydroxytryptamine (5‐HT, serotonin) acts as a neurotransmitter within the central nervous system (CNS) and peripheral nervous system, in addition to mediating nonneuronal actions in numerous other tissues including the gastrointestinal tract and blood vessels. The actions of 5‐HT are, therefore, considered to be copious and diverse. With respect to its role as a neurotransmitter in the brain, which will be the primary focus of this chapter, 5‐HT is implicated in the processes of mood, sleep, aggression, cognition, memory, emesis, and feeding behavior, as well as the pathophysiology of disorders including major depression, schizophrenia, obsessive–compulsive disorder, and anxiety. The impact of this plethora of roles has consequently led to the actions of 5‐HT being extensively studied, leading to the development of many compounds of therapeutic value, including antidepressants, antipsychotics, and antiemetics. This chapter begins with an outline of the 5‐hydroxytryptaminergic system in the brain, including the origins and projections of 5‐HT‐containing neurons, subsequently focusing on the roles of the individual 5‐HT receptors (> Table 8-1) and the transporter in both physiological and pathophysiological processes.

2

Neuroanatomy of the 5‐Hydroxytryptaminergic System

The cell bodies of 5‐HT neurons are situated along the rostrocaudal midline of the brain stem, as first identified by Dahlstro¨m and Fuxe in 1964 (Dahlstro¨m and Fuxe, 1964; see Hornung, 2003, for an in‐depth review), and are categorized into nine anatomical groups, named B1–9. The dorsal raphe nucleus (B6, B7), median raphe nucleus (B8), and B9 contain
85% of the 5‐HT neurons found within the brain, and project extensively to widespread regions of the forebrain. The remaining raphe nuclei, B1–4, innervate primarily both the brain stem and spinal cord. Although the raphe nuclei contain predominantly 5‐HT neurons, it should be noted that other neuronal phenotypes are also apparent.

3

Forebrain Projections of the Raphe Nuclei

Raphe nuclei neurons project to widespread regions throughout the cerebral hemispheres (see Hensler, 2006, for review) via two main pathways, the dorsal periventricular pathway and the ventral tegmental radiation, which unite in the hypothalamus before continuing along the medial forebrain bundle. The hypothalamus, medial septum, and dorsal hippocampus receive predominant innervation from the median raphe nucleus, whereas the ventral hippocampus, amygdala, lateral septum, striatum, and prefrontal cortex (PFC) contain 5‐HT neuron terminals largely from the dorsal raphe nucleus. The cerebral cortex receives inputs from both subdivisions, though various regions are thought to receive different degrees of median and dorsal raphe neuron innervation.

4

The Physiology of 5‐HT Neurons

5‐HT neurons have distinctive electrophysiological properties (e.g., Aghajanian and Haigler, 1974), confirmed using a combination of electrophysiology and fluorescent immunohistochemistry, enabling the activity of 5‐HT‐immunopositive cells to be recorded (Beck et al., 2004). 5‐HT neurons appear to fire

8

5‐Hydroxytryptamine in the central nervous system . Table 8-1 Summary of the structure, pharmacology, and function of 5‐HT receptors Receptor Human gene

5‐HT1A 5q11.2–q13

5‐HT1B 6q13

5‐HT1D 1p34.3–36.3

Structure Transduction system

GPCR ↓cAMP G‐protein‐coupled‐Kþ current 8‐OH‐DPAT

GPCR ↓cAMP

GPCR ↓cAMP

Sumatriptan

Sumatriptan

(R)‐UH301

L 694247

PNU 109291

U92016A WAY 100635 (S)‐UH301 NAD299 (robalzotan) ↑Acetylcholine

GR 55562 SB 224289 SB 236057 ↓5‐HT

Agonists

5‐ht1E 6q14– q15 GPCR ↓cAMP

5‐HT1F 3q11 GPCR ↓cAMP

LY 344864 –

Antagonists

Effect on neurotransmission

Noradrenaline ↓Dopamine Therapeutic target

Depression Anxiety/stress/panic Aggression Cognition

L 694247

↑Acetylcholine ↑Glutamate ↓Dopamine Depression Anxiety Aggression Migraine Drug addiction

Receptor

5‐HT2A

5‐HT2B

5‐HT2C

Human gene

13q14–q21

2q36.3–2q37.1

Xq24

Structure Transduction system

GPCR ↑PLC

GPCR ↑PLC

Agonists

DOI

Antagonists

Ketanserin MDL 100907

GPCR ↑PLC DOI BW 723C86 Ro 600175 RS 127445 SB 200646 SB 204741

Effect on neurotransmission

↑Glutamate ↑Dopamine

Therapeutic target

Depression Anxiety Schizophrenia Cognition Eating disorders Sleep disorder?

LY 334370

BRL 15572 SB 714786

–

–

↑Glutamate

–

–

Migraine

–

Migraine

DOI Ro 600175 SB 242084 RS 102221

? ↓Dopamine

Depression Anxiety Sleep disorder? Migraine

Anxiety Obesity Cognition

5‐HT3 11q23.1–23.2 (A) 11q23.1 (B) 3q27 (C/D/E) LGIC Ion conductance (Kþ, Naþ, Ca2þ) 2‐Methyl 5‐HT SR 57227 m‐Chlorophenyl biguanide DOI Granisetron Ondansetron Tropisetron ↑5‐HT ↑Dopamine ↓Acetylcholine Emesis Anxiety Cognition Drug addiction Analgesia Chronic fatigue syndrome

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. Table 8-1 (Continued) Receptor Human gene

5‐HT4 5q31–q33

5‐ht5a 7q36

Structure

GPCR

Transduction system

↑cAMP

Agonists

Antagonists

Effect on neurotransmission

BIMU 8 RS 67506 ML 10302 GR 113808 SB 204070

5‐HT7 10q21–q24

GPCR

GPCR

Not known

↑cAMP

↑cAMP

5‐CT

5‐CT

–

8‐OH‐DPAT

? SB 699551‐ A

–

Ro 630563 SB 271046

SB 258719 SB 269970

SB 357134 ↓Acetylcholine

SB 656104

↑Dopamine ↓Glutamate Cognition Schizophrenia Depression

? ↑ ↓5‐HT

RS 100235 ↓Acetylcholine ↑Dopamine ↑5‐HT

Therapeutic target

5‐HT6 1p35–36

GPCR ↑cAMP ? Ca2þ mobilization ? Kþ current

5‐ht5B 2q11–13 (nonfunctional) GPCR

Cognition

Not known

Not known

Anxiety

Not known

Not known

Anxiety/stress Epilepsy

Depression Schizophrenia Sleep disorder Epilepsy Cognition

spontaneously with a slow, rhythmic activity, and exhibit a relatively long action potential in addition to a large afterhyperpolarization potential (AHP). The rhythmic activity is thought to be generated by a pacemaker cycle mediated by a calcium‐dependent potassium current (Aghajanian, 1990).

5

The 5‐HT Receptors

The ability of 5‐HT to mediate a diverse array of actions can be accounted for by the existence of an imposing number of 5‐HT receptors (Barnes and Sharp, 1999). Numerous 5‐HT receptor families and subtypes have been identified, particularly within the last two decades following the development of techniques in the field of molecular biology. Presently, there are 18 genes that give rise to 14 distinct mammalian 5‐HT receptor subtypes, divided into 7 families, the majority of which are members of the G‐protein‐coupled receptor (GPCR) superfamily, the sole exception being the 5‐HT3 receptor, a ligand‐ gated ion channel. Further, receptor multiplicity is generated through RNA editing (the 5‐HT2C receptor), alternative splicing (5‐HT3, 5‐HT4, and 5‐HT7 receptors), and the putative formation of homo‐ and heterodimers (5‐HT4 and the b2‐adrenoceptor; Berthouze et al., 2005).

5.1 The 5‐HT1 Receptor Family The 5‐HT1 receptor family contains five subtypes, 5‐HT1A, 5‐HT1B, 5‐HT1D, 5‐HT1E, and 5‐HT1F, each having distinct, but overlapping patterns of expression within the brain. This family can be characterized by its inhibitory effect on cellular cAMP levels, although the 5‐HT1A receptor can also activate a G‐protein‐ activated potassium channel independent of second‐messenger cascades involving cAMP.

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5.1.1 The 5‐HT1A Receptor In 1981, Pedigo et al. (Pedigo et al., 1981) identified the 5‐HT1A receptor‐binding site in rat brain, but the sequence encoding the receptor was not isolated until 1987 (Kobilka et al., 1987). The 5‐HT1A receptor inhibits adenylate cyclase activity through coupling to Gi/o proteins, which has been demonstrated both in native tissue (e.g., rat hippocampus) and recombinant cell systems (see Albert et al., 1996, for review). Such coupling has not, however, been observed in the raphe nuclei (Clarke et al., 1996), where receptor activation induces G‐protein‐modulated potassium current which is cAMP‐independent (Aghajanian, 1995). Expression of the 5-HT7 receptor within the brain region thought to control circadian rhythm, the suprachiasmatic nucleus (SCN), and the overlapping pharmacologies of the 5‐HT1A and 5‐HT7 receptors (e.g. activation by 8-OH-DPAT), complicates interpretation of results in this area of research. More recently, immunohistochemical studies using selective 5‐HT1A receptor antibodies have provided greater resolution of receptor expression through light and electron microscopy. Within the raphe nuclei, the 5‐HT1A receptor appears to be expressed somatodendritically by serotonergic neurons projecting to the forebrain, with dendritic receptors predominantly located in extrasynaptic regions (Kia et al., 1996b; Riad et al., 2000; > Figure 8-1a). The receptor is also found in many regions of the forebrain, including the frontal, piriform, and entorhinal cortices, the hippocampus, preoptic areas, lateral and medial septum, the diagonal band of Broca, hypothalamus, amygdala, and thalamic regions (Aznar et al., 2003). Within the isocortex, the receptor is expressed throughout all laminae (with the exception of layer I), where both glutamatergic pyramidal neurons and calbindin‐ and parvalbumin‐positive inhibitory g‐amino butyric acid (GABA)ergic interneurons express the receptor (Aznar et al., 2003). Within the hippocampus, granule and pyramidal cells are also believed to express the 5‐HT1A receptor on both the soma and dendrites (Riad et al., 2000; Aznar et al., 2003; > Figure 8-1b), particularly in the postsynaptic membrane, but also in nonsynaptic regions, of dendritic spines (Kia et al., 1996b). In the medial septum and diagonal band of Broca, the receptor appears to be expressed somatodendritically by cholinergic neurons (Kia et al., 1996a) and by inhibitory interneurons (Aznar et al., 2003). Given the apparent common subcellular location of the 5‐HT1A receptor, it has been suggested that a structural component of the protein may be responsible for targeting the receptor to a somatodendritic location (Darmon et al., 1998). Activation of the somatodendritic 5‐HT1A autoreceptor in the raphe nuclei induces membrane hyperpolarization, leading to reduced 5‐HT neuron excitability, firing, and ultimately a reduction in 5‐HT release in the raphe forebrain projection areas (Aghajanian, 1995; Sharp et al., 1996). 5‐HT1A receptor agonists also inhibit neuronal firing in forebrain regions, including the hippocampus (e.g., Sprouse and Aghajanian, 1988). The release of other neurotransmitters, including acetylcholine, noradrenaline, and dopamine, is thought to be regulated by 5‐HT1A receptor activation. For example, 8‐OH‐DPAT augments acetylcholine release in the hippocampus and cortex of guinea pigs (Bianchi et al., 1990; Wilkinson et al., 1994). The mechanism of this action is unclear, though more recent studies suggest that activation of noncortical presynaptic 5‐HT1A autoreceptors mediate the increase in acetylcholine release within the cortex (Millan et al., 2004), while locally administered 8‐OH‐DPAT‐induced elevation of acetylcholine levels in the rat dorsal hippocampus may be mediated by postsynaptic 5‐HT1A receptors (Nakai et al., 1998). In contrast, other reports suggest that 5‐HT1A receptor antagonists enhance acetylcholine release within the hippocampus by blocking tonically active inhibitory 5‐HT1A receptors on cholinergic cells (Millan et al., 2004), whereas noradrenaline levels in the hippocampus, ventral tegmental area (VTA), and hypothalamus increase following 5‐HT1A receptor activation (Done and Sharp, 1994; Chen and Reith, 1995; Suzuki et al., 1995). More recently, putative 5‐HT1A receptor‐mediated effects on dopamine release have been observed; whereby the 5‐HT1A receptor agonist BAYx3702 elevated dopamine release in both the VTA and medial PFC (mPFC), as well as increasing dopaminergic neuron activity in the VTA (Dı´az‐Mataix et al., 2005). 5‐HT1A receptors may also modulate glutamatergic neurotransmission. Indeed, 5‐HT1A receptor activation attenuates AMPA currents in pyramidal neurons of the PFC, potentially through a reduction in PKA‐dependent AMPA subunit phosphorylation (Cai et al., 2002). The activity of 5‐HT1A receptors may be involved in the pathogenesis and treatment of psychiatric disorders. One such condition that attracted much study is depression. It has been suggested that levels of 5‐HT1A receptors are altered in depressed subjects, although to date, studies to this effect remain

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. Figure 8-1 (a) Low‐power electron micrograph illustrating the somatodendritic localization of the 5‐HT1A immunoreactivity (immunoperoxidase labeling) in the NRD. Most of the field is occupied by the multipolar cell body of a presumed 5‐HT neuron (N1 in nucleus) showing strong immunolabeling of its perikaryon and emerging proximal dendrite (D). Smaller adjacent nerve cell bodies (N2, N3 in nuclei) are immunonegative. Numerous transversely sectioned dendritic branches of smaller calibre (d) are also immunoreactive in the surrounding neurophil, but myelinated axons and axon terminals are all immunonegative. Also note the immunonegativity of a nearby astrocyte (A), pericyte (P), and endothelial cell (E). In the labeled cell body and the largest of the dendritic branches, the diffusible immunoperoxidase precipitate is conspicuously concentrated on the inner face of the plasma membrane, as a rim almost 1 mm wide; in smaller dendritic processes, the confluence of these peripheral zones of 5‐HT1A immunoreactivity accounts for labeling of the entire sectional surface. This and the following electron micrograph were obtained from tissue fixed with acrolein plus paraformaldehyde. Scale bar ¼ 5 mm. (b) Immunoperoxidase labeling of several immunoreactive dendrites (d) in a small field from the stratum radiatum of CA3. The peroxidase immunoprecipitate tends to cluster near the plasma membrane and is also found in dendritic spines (arrows) seen to emerge from their parent dendritic branches in the upper left and lower right corners of the figure. Scale bar ¼ 1 mm (Taken from Riad et al., 2006, these figures are reproduced with permission from the authors)

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controversial. Some reports have suggested that the levels of the receptor in the hippocampus are reduced (Cheetham et al., 1990), whereas others detect no differences between suicide victims suffering from depression and control patients (Stockmeier et al., 1997). Alternatively, abnormally high levels of 5‐HT1A receptors were found in the PFC (Arango et al., 1995) and dorsal raphe nucleus (Stockmeier et al., 1998) of depressed suicide victims, whereas conversely other studies found no change in the cortex (e.g., Lowther et al., 1997) or reductions in the number of 5‐HT1A receptors in the dorsal raphe nucleus (Arango et al., 2001). Receptor levels aside, it has become apparent that compounds acting on the 5‐HT1A receptor may have therapeutic value in treating depression. The use of 5‐HT1A receptor antagonists in conjunction with selective serotonin reuptake inhibitors (SSRIs) has been suggested to reduce the onset of relief from depression (Artigas et al., 1994), although other reports have found no such benefits (Segrave and Nathan, 2005). The 5‐HT1A receptor has also been linked to depression through the ability of the 5‐HT1A receptor agonist, 8‐OH‐DPAT, following 5‐HT depletion, to induce granule cell proliferation within the dentate gyrus of the hippocampus (Huang and Herbert, 2005), a process thought to facilitate the treatment of depression (Malberg et al., 2000; Santarelli et al., 2003). Similarly, the 5‐HT1A receptor has been linked to suicidal behavior in addition to depression. Pitchot et al. (2005) have found that flesinoxan, a 5‐HT1A receptor agonist, attenuated cortisol and temperature responses in suicidal patients when compared with controls, but not in merely depressed patients, suggesting that reduced 5‐HT1A receptor sensitivity may be connected to suicidal tendencies. Initial reports using a 5‐HT1A receptor knockout mouse strain demonstrated various behavioral abnormalities compared with wild‐type mice, including increased levels of anxiety in the open‐field arena test and attenuated immobility in the forced‐swim test. This latter effect may represent either increased anxiety due to the inherent stress of the testing or an inhibition of behavioral despair, a measure of predisposition to depression (Parks et al., 1998; Ramboz et al., 1998). These observations correlate with the observation that 5‐HT1A receptor agonists are anxiolytic (Lucki et al., 1994). Indeed, buspirone, a partial 5‐HT1A receptor agonist, has therapeutic usage in the treatment of generalized anxiety disorder (Kapczinski et al., 2003), while exhibiting minimal side effects relative to those induced following administration of benzodiazepines for anxiety‐related disorders (Goa and Ward, 1986). Recently, new 5‐HT1A receptor ligands (oMPP derivatives), supposed postsynaptic antagonists and partial agonists, display anxiolytic activity when assessed in the Vogel conflict drinking test (Bojarski et al., 2006). 5‐HT1A receptor targeting, therefore, continues to provide a useful strategy in the search for novel treatments of anxiety and depression. In addition to anxiety, the 5‐HT1A receptor may also be involved in responses to chronic stress implicated in depression. It has been shown that chronic stress may lead to the downregulation of hippocampal 5‐HT1A receptors, potentially through the actions of glucocorticoids in suppressing gene transcription (Wissink et al., 2000). In addition, glucocorticoids may affect 5‐HT1A receptor sensitivity, as loss of glucocorticoids by adrenalectomy resulted in a leftward shift of the 5‐HT1A receptor concentration– response curve in the CA3 region of the hippocampus (Okuhara and Beck, 1998). The 5‐HT1A receptor is also thought to be involved in panic responses generated by the dorsal raphe nucleus–dorsal periaqueductal gray pathway. Pobbe and Zangrossi (2005), for example, demonstrated that injection of the 5‐HT1A receptor antagonist, WAY 100635, into the dorsal raphe nucleus impaired escape in the elevated T‐maze, indicating a reduced level of panic, presumably resulting from the inhibition of tonic 5‐HT1A autoreceptor activity. 5‐HT1A receptor activity may be involved in mediating aggressive behavior. Agonists at the 5‐HT1A receptor, for example, have been shown to inhibit aggressive behavior in rodents and humans, though they also sedate, which complicates interpretation (de Boer and Koolhaas, 2005). A recent study, however, used an atypical 5‐HT1A receptor ligand, S‐15535, which acts as an agonist at the somatodendritic 5‐HT1A receptor, while acting as an antagonist, or weak partial agonist, at postsynaptic receptors (de Boer and Koolhaas, 2005). This compound exhibited antiaggressive properties, suggesting that 5‐HT1A receptor agonism at the presynaptic level is responsible for the antiaggressive effects. Furthermore, administration of both S‐15535 and alnespirone (the latter being agonist at both pre‐ and postsynaptic 5‐HT1A receptors) showed an additive effect in reducing aggression rather than a predicted attenuation of the alnespirone‐ mediated effect if the actions of S‐15535 were postsynaptic.

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The control of rodent circadian rhythm may also be affected by 5‐HT1A receptor activation. Weber et al. (1998) demonstrated that compounds with 5‐HT1A receptor agonist activity were able to suppress light‐ induced phase shifts in the hamster circadian rhythm. In addition, Gannon and Millan (2006) showed that S‐15535 increased hamster light‐induced phase advances. Investigation of the 5‐HT1A receptor knockout mouse also discovered the potential role of this receptor subtype in sleep, specifically with respect to the paradoxical sleep phase. The knockout strain showed signs of increased periods of paradoxical sleep, while WAY 100635 induced similar effects in the wild‐type mouse (Boutrel et al., 2002). Expression of the 5‐HT7 receptor within the brain region thought to control circadian rhythm and the suprachiasmatic nucleus (SCN); however, the overlapping pharmacologies of the 5‐HT1A and 5‐HT7 receptors (e.g., activation by 8‐OH‐DPAT) has complicated this area of research. Studies with the 5‐HT1A receptor knockout mouse have led to suggestions that this subtype may take part in the processes of memory and learning. Sarnyai et al. (2000) investigated the performance of knockout mice in the Morris water maze and Y‐maze tests, and found knockout mice had lower levels of performance compared with wild‐type subjects, suggesting that the 5‐HT1A receptor has a positive effect on hippocampal‐dependent learning and memory. In contrast, studies with the 5‐HT1A receptor agonist 8‐OH‐DPAT suggest that activation of the receptor impairs working and spatial memory function. For example, systemic administration of 8‐OH‐DPAT impaired performance of rats in the radial maze, an effect blocked by the 5‐HT1A receptor antagonist, WAY 100635 (Helsley et al., 1998; Egashira et al., 2006). Egashira and colleagues further suggest that this effect of 8‐OH‐DPAT is mediated by postsynaptic 5‐HT1A receptors in the dorsal hippocampus as the effects of systemic administration were mimicked by local injection into the dorsal hippocampus, but not into other regions. It is possible, however, that 8‐OH‐ DPAT induces a biphasic response, as high doses impair memory and low doses have been shown to attenuate scopolamine‐ and tetrahydrocannabinol‐induced memory impairment, potentially through enhancing acetylcholine release in the dorsal hippocampus (Inui et al., 2004). Further evidence for 5‐HT1A receptor activation impairing cognitive processes has been provided in human studies, where administration of tandospirone, a 5‐HT1A receptor agonist, inhibited verbal memory (Yasuno et al., 2003), and psilocybin, a 5‐HT1A/2A receptor agonist, reduced performance in attentional tracking, but not in spatial working memory, in the presence of the 5‐HT2A receptor antagonist ketanserin (Carter et al., 2005). One recent development in our understanding of the role of the 5‐HT1A receptor in neurological disorders is the possible involvement of this subtype in Alzheimer’s disease. A recent positron emission tomography (PET) study of Alzheimer’s patients demonstrated reduced 5‐HT1A receptor‐binding sites in both the hippocampus and raphe nuclei, having accounted for a loss in neuronal volume, which was correlated with the severity of the observed clinical symptoms (Kepe et al., 2006). If future studies confirm the 5‐HT1A receptor is involved, then ligands at this receptor subtype may have therapeutic benefit in treating Alzheimer’s disease.

5.1.2 The 5‐HT1B Receptor The 5‐HT1B receptor‐binding site was initially distinguished from the 5‐HT1A receptor due to its low affinity for 8‐OH‐DPAT (Middlemiss and Fozard, 1983) and the rat receptor sequence was identified in 1991 by Voigt et al. (Voigt et al., 1991). During the cloning of the numerous 5‐HT1 receptor genes, there was some confusion as to whether newly identified rodent and human sequences simply represented species differences of the same subtype, or if they encoded distinct receptor subtypes. For instance, initially the human 5‐HT1B receptor sequence was classified as 5‐HT1Db, though it has now been reclassified (Hartig et al., 1996). The distribution of the 5‐HT1B receptor in the brain has been extensively characterized through receptor autoradiography (Pazos and Palacios, 1985) and immunohistochemistry (Sari et al., 1997, 1999), whereas the location of 5‐HT1B receptor mRNA has been determined by in situ hybridization (Boschert et al., 1994; Varnas et al., 2005). Given the similarity in pharmacological profile with the 5‐HT1D receptor, however, early studies reported that the detection of the 5‐HT1B receptor‐binding site may have been unable to distinguish between the two subtypes (see later). The 5‐HT1B receptor appears to

5‐Hydroxytryptamine in the central nervous system

8

be expressed at highest levels in the basal ganglia, particularly the globus pallidus and substantia nigra, with lower levels being found in the periaqueductal gray, superficial layer of the superior colliculus, cortex, amygdala, hypothalamus, hippocampus, cerebellum, and dorsal horn of the spinal cord (Bonaventure et al., 1997; Sari et al., 1999; Varnas et al., 2001). A more detailed study of the ultrastructural receptor location suggests a predominant location on axons and axon terminals, though not on synaptic membranes, as has been demonstrated within the substantia nigra and globus pallidus (Sari et al., 1997, 1999). Correspondingly, the distribution of receptor transcripts does not completely match the location of the receptor‐ binding sites, as 5‐HT1B mRNA has been identified in the raphe nuclei, striatum, hippocampus, cortex, and thalamus (Varnas et al., 2005). Transcripts are notably absent from the substantia nigra and globus pallidus, which display the strongest levels of binding sites. The 5‐HT1B receptor is thought to act as an auto‐ and heteroreceptor on 5‐HT and non‐5‐HT neurons, respectively. 5‐HT1B receptor activation has been shown to mediate inhibition of 5‐HT release in the forebrain, including the frontal cortex and hippocampus (Sleight et al., 1989; Bosker et al., 1995; Trillat et al., 1997). In addition, studies with the 5‐HT1B knockout mouse detected lower levels of 5‐HT within the nucleus accumbens and locus coeruleus (Ase et al., 2000). With respect to an impact on other neurotransmitter systems, the 5‐HT1B receptor inhibits acetylcholine release in the hippocampus from septal afferents (Cassel et al., 1995), suppresses glutamatergic transmission in the subiculum (Sari, 2004), and inhibits GABA release in the substantia nigra (Stanford and Lacey, 1996). The 5‐HT1B receptor knockout mouse also showed reduced basal dopamine levels in the nucleus accumbens, indicative of increased dopamine turnover (Ase et al., 2000). The 5‐HT1B receptor has been suggested to play a role in many physiological and pathophysiological processes. Firstly, this receptor subtype may contribute to the generation of anxious states; selective 5‐HT1B receptor agonists have been demonstrated to be anxiogenic (Lin and Parsons, 2002), potentially acting through receptors in the hippocampus and amygdala. In support of these findings, it has been shown that overexpression of the 5‐HT1B receptor in the dorsal raphe nucleus elevates stress‐induced anxiety (Clark et al., 2002). The 5‐HT1B receptor may also have a role in the generation or treatment of depression, illustrated by potential interactions between antidepressant treatment and 5‐HT1B receptor function. Chronic treatment with SSRIs, for example, downregulates or desensitizes 5‐HT1B receptors in the SCN (O’Connor and Kruk, 1994), and reversibly reduces levels of 5‐HT1B receptor transcripts in the dorsal raphe nucleus (Anthony et al., 2000), while receptor antagonism augments SSRI‐induced increases in frontal cortex 5‐HT levels (Gobert et al., 1997). In addition, a behavioral study has demonstrated that 5‐HT1B receptor activation reduces immobility in the mouse forced‐swim test, most likely via heteroreceptors, which forward a therapeutic strategy for the treatment of depression (Tatarczynska et al., 2005). Further supporting evidence comes from a recent study by Svenningsson et al. (2006). Using the yeast‐two‐hybrid system, they identified a molecular interaction between the 5‐HT1B receptor and p11, an S100 EF‐hand protein. Interestingly, p11 expression is reduced in both animal models of depression and in the human brains of depressed patients, whereas expression is upregulated on pharmacological (e.g., imipramine) or electroconvulsant therapy (> Figure 8-2). Furthermore, p11 knockout mice display symptoms of depression. It appears that p11 has a role in targeting the 5‐HT1B receptor to the cell surface, as well as facilitating receptor transduction. It is possible, therefore, that loss of 5‐HT1B receptor function through defective p11 expression may contribute to the pathogenesis of depression. Like the 5‐HT1A receptor, it has been suggested that the 5‐HT1B receptor is involved in regulating aggressive behavior. The 5‐HT1B receptor knockout mouse shows signs of increased aggression in resident– intruder paradigms (Saudou et al., 1994). Likewise, 5‐HT1B agonists, including CP 93129 and CGS 12066B, appear to reduce levels of aggression (de Boer and Koolhaas, 2005). It is unclear, however, whether these effects are mediated by presynaptic auto‐ or heteroreceptors (de Boer and Koolhaas, 2005), although the anterior hypothalamus has been forwarded as a putative location for serotonergic modulation of aggressive behavior (Ferris et al., 1997). The putative ability of the 5‐HT1B receptor to modulate responses to addictive drugs has also great therapeutic potential. For example, 5‐HT1B receptor activation reduces the self‐administration of alcohol in rats (Tomkins and O’Neill, 2000) and 5‐HT1B receptor knockout mice consume more alcohol (Crabbe et al., 1996). This 5‐HT1B receptor activity may be mediated by its regulation of dopaminergic neurotransmission,

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. Figure 8-2 Regulation of p11 expression by antidepressant treatments and in depression‐like states. In situ hybridization illustrating an upregulation of p11 mRNA in the forebrain following (a) repeated treatment with imipramine [10 mg/kg per day, intraperitoneally (i.p.) for 14 days] in mice (n ¼ 8 per group) and (b) electroconvulsive therapy (ECT) for 10 days in rats (n ¼ 5 per group). Conversely, p11 mRNA was downregulated in (c) the forebrain in helpless H/Rouen versus nonhelpless NH/Rouen mice (n ¼ 10 per group), and (d) in patients who suffered from unipolar major depression (n ¼ 15 per group). Data from the anterior (a; b; c, left; d) and posterior (c, right) cingulated cortices were normalized to the corresponding controls and represent means  SEM. *p < 0.05, ***p < 0.001 versus control by student’s t test (This figure is reproduced with permission from the authors (Svenningsson et al., 2006))

as demonstrated by Yan et al. (2005) by showing that ethanol‐induced increases in dopamine neuron activity in the VTA were suppressed by the selective 5‐HT1B receptor antagonist, SB 216641, and enhanced by the 5‐HT1B receptor agonist, CP 94253. Evidence has also been presented for the involvement of the 5‐ HT1B receptor in the reinforcing properties of cocaine. Studies using the 5‐HT1B receptor knockout mouse demonstrated elevated dopamine levels in the nucleus accumbens, both basally and following cocaine administration (Shippenberg et al., 2000).

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Modulation of central dopamine function may also be relevant for the 5‐HT1B receptor’s role in locomotor activity demonstrated by the ability of the 5‐HT1B receptor agonist, RU 24969, to increase spontaneous movement and the increased exploratory behavior in 5‐HT1B receptor knockout mice (Ramboz et al., 1996; Brunner et al., 1999). The circadian rhythm in rodents may also be modulated by the actions of the 5‐HT1B receptor. The 5‐HT1B receptor has been identified within the SCN of the hypothalamus, a key region involved in controlling circadian rhythm. In 2000, Garabette et al. (Garabette et al., 2000) demonstrated that local administration of the 5‐HT1B receptor agonist, RU 24969 attenuated 5‐HT release in the SCN. In addition, 5‐HT1B receptor agonists block glutamatergic excitatory postsynaptic currents (EPSCs) within this region (Smith et al., 2001), and reduce the frequency of inhibitory postsynaptic currents (IPSCs) in GABAergic SCN neurons (Bramley et al., 2005). Study of the 5‐HT1B receptor knockout mouse identified an inability of this strain to entrain to unnatural photic stimuli (i.e., those not based on a 24‐h cycle), suggesting that the absence of this receptor subtype affects the ability of the circadian rhythm to respond to changes in light (Sollars et al., 2006). At present, perhaps the most important therapeutic application of 5‐HT1B receptor ligands involves the treatment of migraine, which is both a neural and vascular disorder. The receptor inhibits trigeminal sensory nerve activity with a subsequent reduction in relevant neuropeptide release. In addition, the 5‐HT1B receptor is expressed by the endothelium of meningeal vessels, which undergo an inflammatory response during migraine, and hence regulate the contractile activity of these vessels. The strategic development of 5‐HT1B receptor agonists (e.g., sumatriptan, zolmitriptan, and naratriptan) as antimigraine agents has been vindicated and made an important impact on the treatment of migraine. Unlike other 5‐HT receptors, the 5‐HT1B receptor appears to be modulated by an endogenous tetrapeptide (Leu–Ser–Ala–Leu), named 5‐HT moduline. This peptide, first isolated from rat and bovine brain in 1996 (Rousselle et al., 1996), appears to inhibit 5‐HT binding to the receptor through an allosteric site distinct from the ligand‐binding domain, and can be released, Ca2þ/Kþ dependently, from rat cortical synaptosomes (Massot et al., 1996). Furthermore, there appears to be a functional consequence to this interaction since 5‐HT moduline has a 5‐HT1B receptor antagonist‐like effect in the mouse social‐ interaction test and the actions of the 5‐HT1B receptor agonist, RU 24969, are inhibited (Massot et al., 1996). In addition, presumed block of 5‐HT moduline action with an antibody reduced measures of anxiety in the open‐field test (Grimaldi et al., 1999). The receptor interaction of this endogenous peptide may, therefore, offer a therapeutic strategy for the treatment of anxiety, depression, and aggressive behavior.

5.1.3 The 5‐HT1D Receptor The human 5‐HT1D receptor (formally known as the 5‐HT1Da receptor) was initially cloned in 1991 by Hamblin and Metcalf (Hamblin and Metcalf, 1991). The 5‐HT1D receptor is relatively less well understood in comparison with the 5‐HT1A and 5‐HT1B receptors, primarily due to the difficulties in separating 5‐HT1B and 5‐HT1D receptors pharmacologically. Consequently, selective identification of 5‐HT1D receptor‐binding sites has been difficult. Receptor autoradiography using, for instance, [125I]GTI with CP 93129 to mask 5‐HT1B receptor sites, identified 5‐HT1D‐like binding sites in the rat basal ganglia (globus pallidus, substantia nigra, and caudate putamen), and the hippocampus and cortex (Bruinvels et al., 1993). Studies with human brain tissue exploited the 15–30‐fold greater selectivity of ketanserin for the 5‐HT1D receptor and determined ketanserin‐sensitive [3H]sumatriptan‐binding sites (Castro et al., 1997), where the distribution of 5‐HT1D‐like sites was comparable with rat brain. In situ hybridization studies in rat and primate brain localized transcripts to the dorsal raphe nucleus, locus coeruleus, nucleus accumbens, olfactory cortex, and caudate putamen (Bruinvels et al., 1994). Similar to the 5‐HT1B receptor, mRNA was not identified in the globus pallidus and substantia nigra, suggesting that the receptor protein may be found on axon terminals projecting from other regions (caudate putamen?). The 5‐HT1D receptor inhibits adenylate activity in recombinant cell systems (Hamblin and Metcalf, 1991). With respect to physiological functions, the nature of receptor location suggests that it could act as an auto‐ or heteroreceptor. It is generally accepted, however, that at least the principle terminal 5‐HT

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autoreceptor, in terms of function, is the 5‐HT1B receptor due to sensitivity to the relatively selective 5‐HT1B receptor antagonist, SB 216641, but not the 5‐HT1D receptor antagonist, BRL 15572 (Schlicker et al., 1997). However, some evidence for a heteroreceptor function is available. For instance, Maura et al. (1998) demonstrated a proposed 5‐HT1D receptor‐mediated inhibition of glutamate release in the human cortex. The performance of this study, however, predated the availability of selective ligands and hence a definitive central role for the 5‐HT1D receptor remains to be determined. This 5‐HT receptor subtype may play a role in the treatment of migraine; however, the efficacious antimigraine ‘‘triptans’’ are 5‐HT1D receptor agonists (as well as 5‐HT1B receptor agonists). To further support this premise, the 5‐HT1D receptor is expressed in the trigeminal ganglia (Hou et al., 2001). The recent discovery of a highly selective 5‐HT1D receptor antagonist, SB 714786, should aid the investigation of the physiological functions of the 5‐HT1D receptor (Ward et al., 2005).

5.1.4 The 5‐ht1E Receptor The 5‐ht1E receptor remains the only member of the 5‐HT1 receptor family that retains the lower class appellation of a protein with unidentified function. The human gene was isolated in 1992 (McAllister et al., 1992), but there was a subsequent lack of rodent 5‐ht1E cloned sequences. Recently, however, the guinea pig 5‐ht1E receptor gene was identified, whereas the mouse genome has been found to contain no 5‐ht1E receptor sequence (Bai et al., 2004). 5‐ht1E receptor distribution within the brain has been documented, with transcripts found in the cortex, caudate putamen, and amygdala of human tissue (Bruinvels et al., 1994). Attempts to map the receptor‐binding sites have exploited the low affinity of the 5‐ht1E receptor for 5‐CT to distinguish it from most other 5‐HT1 receptors (Miller and Teitler, 1992). Putative 5‐ht1E receptor‐binding sites were found in the cerebral cortex, caudate putamen, and claustrum, with lower levels of binding in the hippocampus and amygdala. The apparent colocalization of mRNA and receptor suggests a postsynaptic distribution for this receptor subtype. Similar to other members of the 5‐HT1 family, within recombinant cell systems the 5‐ht1E receptor couples negatively to adenylate cyclase (Levy et al., 1992). Study of this potential receptor is hindered by the lack of selective ligands, although structure–affinity relationship (SAR) studies have been reported (e.g., Dukat et al., 2004), which demonstrate a similar pharmacology to the 5‐HT1F receptor, although a few notable distinctions are apparent (see later). Hence, such studies indicate that selective ligands for the 5‐ht1E receptor may be identifiable.

5.1.5 The 5‐HT1F Receptor The 5‐HT1F receptor was initially identified in the mouse genome, but was named 5‐ht1Eb due to its similar pharmacological to the 5‐ht1E receptor (Amlaiky et al., 1992), though localization of 5‐HT1F receptor transcripts clearly showed a distinct pattern of expression when compared with the 5‐ht1E receptor. Receptor autoradiography with [3H]sumatriptan in the presence of 5‐CT to block 5‐HT1B/1D receptor‐binding sites demonstrates that the 5‐HT1F receptor is located within the guinea pig hippocampus, cortex, claustrum, and caudate nucleus (Waeber and Moskowitz, 1995a, b). More recently, the selective ligand [3H]LY 334370 confirmed largely the distribution of the 5‐HT1F receptor in rodent brain (Lucaites et al., 2005). Despite the identification of two 5‐HT1F receptor selective ligands, LY 344864 and LY 334370, little information has been published concerning the physiological function(s) of this receptor. One potential role is an involvement in the treatment of migraine; forwarded initially due to the relatively high affinity of sumatriptan and naratriptan. In support of this hypothesis, the 5‐HT1F receptor has been localized on glutamatergic neurons within the trigeminal ganglia (Ma, 2001); activation of these neurons are thought to induce dural protein extravasation, a potential contributor to the generation of migraine, and the 5‐HT1F receptor agonist, LY 344864, inhibits this process (Phebus et al., 1997).

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5.2 The 5‐HT2 Receptors The 5‐HT2 receptor family consists of three receptor subtypes: 5‐HT2A, 5‐HT2B, and 5‐HT2C receptors (the latter was previously known as 5‐HT1C receptor, but was appointed to the 5‐HT2 receptor family on the basis of its structure, mode of signal transduction, and to some extent pharmacology; Hoyer et al., 1994). Until fairly recently, it has been difficult to distinguish the three subtypes pharmacologically, but now selective antagonists including MDL 100907 (5‐HT2A), SB 204741 (5‐HT2B), and SB 242084 (5‐HT2C) have become available to allow responses to be assigned to individual receptors with more confidence.

5.2.1 5‐HT2A Receptor The 5‐HT2A receptor was initially identified as a binding site in rat cortical membranes (Peroutka and Snyder, 1979), with subsequent identification of the rat sequence a decade later (Pritchett et al., 1988; Julius et al., 1990). The distribution of the 5‐HT2A receptor in the brain has been well characterized. Receptor autoradiography with selective ligands, such as [3H]‐MDL 100907, has shown high levels of expression in human and rodent forebrain, including the neocortex, entorhinal and piriform cortices, hippocampus, caudate nucleus, nucleus accumbens, and olfactory tubercle (Lopez‐Gimenez et al., 1997). The location of 5‐HT2A mRNA corresponds well to receptor distribution (Burnet et al., 1995), generally following the distribution of 5‐HT neuron innervation, implying that the receptor has a postsynaptic location. The cellular expression of the 5‐HT2A receptor protein appears to be predominantly neuronal, both on GABAergic interneurons in the cortex and glutamatergic pyramidal cells within the cortex and hippocampus (Pompeiano et al., 1994; Burnet et al. 1995; Willins et al., 1997; Jakab and Goldman‐Rakic, 1998). A detailed study of the subcellular location of 5‐HT2A receptors in rat PFC (Miner et al., 2003) observed expression on both the shafts and spines of proximal and distal pyramidal dendrites, reinforcing the likely postsynaptic location of the receptor. In addition to the 5‐HT2A receptor being expressed on the plasma membrane (Willins et al., 1997), it also exhibits a degree of intracellular localization, which may be indicative of a high rate of receptor turnover (Martin‐Ruiz et al., 2001; Cornea‐Hebert et al., 2002). The precise ultrastructural positioning of the receptor is supported by the work of Cornea‐Hebert et al. (2002), who demonstrated that the 5‐HT2A receptor physically interacts with the cytoskeletal protein, MAP1A, suggesting that the receptor may regulate neuronal development or dendritic plasticity. The 5‐HT2A receptor is coupled to the activation of phospholipase C (PLC), inducing the mobilization of intracellular Ca2þ stores, in both recombinant systems and native tissue (Conn and Sanders‐Bush, 1984; Pritchett et al., 1988). Additionally, the 5‐HT2A receptor may activate second‐messenger cascades responsible for the regulation of brain‐derived neurotrophic factor (BDNF) levels, as 5‐HT2A receptor agonists reduce levels of BDNF in the dentate gyrus of the hippocampus, while increasing levels in the neocortex, which has potentially profound effects on neuronal growth (Vaidya et al., 1997). 5‐HT2A receptor activation is also thought to mediate, at least in part, the 5‐HT‐induced attenuation of the slow after hyperpolarizing current observed in layer V pyramidal neurons in the cortex following a burst of spikes (Villalobos et al., 2005). The 5‐HT2A receptor regulates the release of many neurotransmitters, including glutamate, dopamine, and GABA. Within the forebrain, for example, the 5‐HT2A receptor increases both glutamate release from layer V pyramidal neurons in the PFC (see Aghajanian and Marek, 1999b, for review) and GABA release onto CA1 pyramidal neurons in the hippocampus (Shen and Andrade, 1998). The 5‐HT2A receptor also appears to have a regulatory effect on dopaminergic neuron firing, supported by receptor expression being associated with dopaminergic neurons within the VTA and substantia nigra (Ikemoto et al., 2000). Furthermore, 5‐HT2A receptor antagonism attenuates dopamine release in the VTA (De Deurwaerdere and Spampinato, 1999) and striatum (Lucas and Spampinato, 2000). It has been suggested that the 5‐HT2A may be involved in the pathogenesis of depression and mediate some of the effects of antidepressant treatment. Correspondingly, the selective knockout of 5‐HT2A receptor expression through injection of antisense oligonucleotides evokes antidepressant‐like effects in the mouse forced‐swim test (Sibille et al., 1997), and also reduced anxiety in the elevated plus maze paradigm

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(Cohen, 2005). Indeed, it has been suggested that levels of 5‐HT2A receptor expression are altered in the ‘‘depressed’’ brain. One PET study, for example, identified reductions in 5‐HT2A receptor levels within the frontal, occipital, temporal, and cingulate cortices of drug‐naı¨ve depressed patients relative to control subjects (Messa et al., 2003). The evaluation of drug‐naı¨ve depressed patients here is likely to be important. For instance, the many studies assessing 5‐HT2A receptor levels in human postmortem tissue from depressed or suicide victims have generally reported conflicting results, with some papers finding elevated 5‐HT2A receptor levels (e.g., Pandey et al., 2002), whereas others are unable to identify any differences. The complication of therapy impacting on 5‐HT2A receptor expression is difficult to control in these postmortem studies. Relevant to potential alterations in the expression of the 5‐HT2A receptor, studies of the physiological actions of the 5‐HT2A receptor have postulated a therapeutic role in the treatment of depression, as 5‐HT2A receptor activation has a negative feedback on raphe neuron firing (Boothman et al., 2003). It has, therefore, been suggested that antagonism of the 5‐HT2A receptor might accelerate the rate of onset of antidepressant drugs, similar to 5‐HT1A receptor antagonism. Consequently, Boothman et al. (2006) reported an elevation in hippocampal 5‐HT release following administration of the SSRI citalopram in combination with a 5‐HT2A receptor antagonist. The authors postulated that this effect could be due to either a block of terminal 5‐HT2A receptor‐mediated negative feedback to the raphe neurons, or through inhibiting GABAergic transmission by interneurons expressing the 5‐HT2A receptor in the raphe nucleus itself. The 5‐HT2A receptor may also contribute to depression through modification of BDNF levels, which increase as a consequence of antidepressant treatment (Nibuya et al., 1995). Furthermore, expression of BDNF and the 5‐HT2A receptor appear to be linked, as a mutant mouse with low levels of postnatal BDNF expression displayed a significant deficit in 5‐HT2A receptor expression in both the dorsal raphe nucleus and PFC, and exhibited a corresponding reduction in the 5‐HT2A receptor‐mediated postsynaptic response in the mPFC (Rios et al., 2006; > Figure 8-3). The 5‐HT2A receptor is also thought to play an integral role in the treatment of schizophrenia, in addition to mediating the effects of hallucinogenic drugs, both of which may have similar underlying mechanisms (Vollenweider and Geyer, 2001). The most compelling evidence that the 5‐HT2A receptor is involved in the pathogenesis and/or the treatment of schizophrenia has come from the study of atypical antipsychotic pharmacology, as most, but not all, of these compounds have a high affinity for the 5‐HT2A receptor (for review, see Meltzer et al., 2003). The most likely region to generate both of these actions is the . Figure 8-3 5‐HT2A‐mediated postsynaptic responses in the prefrontal cortex. (a) Cumulative probability curves showing that serotonin elicits a significant increase in sEPSC frequency in neurons from wild‐type mice (K‐S test; p < 0.001) but not BDNF‐mutant (p ¼ NS) mice. (b) Bar graph summarizes data (mean  SE) for WT and BDNF‐mutant mice. The data are based on recordings from wild‐type cells (n ¼ 5; two mice) and cells from BDNF2L/2LCamKCse93‐mutant mice (n ¼ 9; four mice); t test, p < 0.001 (This figure is reproduced with permission from the authors (Rios et al., 2006))

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cortex, where glutamatergic pyramidal neurons have been shown to express the 5‐HT2A receptor, and to be regulated by its activity (Aghajanian and Marek, 1999a, b). Furthermore, some studies have shown a reduction in both 5‐HT2A receptor transcripts and binding sites in the cortex and parahippocampal gyrus of schizophrenic patients (Burnet et al., 1996), although a PET study appears to contradict these findings (Okubo et al., 2000). Learning and memory may be affected by 5‐HT2A receptor activation, given the high levels of receptor expression within the hippocampus and cortex, both key regions in mnemonic function. Of more direct evidence for instance, 5‐HT2A receptor agonists have been shown to improve rabbit eyeblink response, a measure of associative learning, though the compounds used are also 5‐HT2C receptor agonists (Welsh et al., 1998; see Harvey, 2003, for review). A role for the 5‐HT2A receptor in working memory has also been postulated (Williams et al., 2002), due to reduced performances of primates in delayed response tasks following administration of selective 5‐HT2A receptor antagonists, while agonists induced a modest improvement in performance. Interestingly, working memory is thought to be deficient in schizophrenia, further implying that this receptor subtype is involved in the pathogenesis of this disorder. Secretion of hormones from the hypothalamus, including ACTH, corticosterone, oxytocin, prolactin, and rennin, has been found to be under the control of 5‐HT, probably via the 5‐HT2A receptor, since the response is mimicked by the 5‐HT2A/2C receptor agonist DOI and antagonized by the selective 5‐HT2A receptor antagonist, MDL 100907 (Van de Kar et al., 2001). Anorexia and bulimia nervosa are psychological conditions thought to involve the 5‐HT system, due to the occurrence of abnormalities in mood, feeding, and impulsiveness, in addition to the ability of drugs acting on the 5‐HT system being only the compounds to date displaying efficacy to treat these conditions. Little other direct evidence has been found, although Audenaert et al. (2003) have shown an apparent reduction in 5‐HT2A receptor binding in the frontal, parietal, and occipital cortices of patients with eating disorders using SPECT brain imaging and the selective ligand, [123I]‐5‐I‐R91150. The 5‐HT2A receptor may also be involved in the regulation of a particular phase of sleep, termed nonrapid eye movement sleep (NREMS). The relatively nonselective 5‐HT2 receptor antagonist, ritanserin, increases NREMS in human subjects (Idzikowski et al., 1991), while blocking the 5‐HT2A receptor in mice has the same effect and this response was not evident in 5‐HT2A receptor knockout strains (Popa et al., 2005). Further investigation of the role of the 5‐HT2A receptor in sleep is clearly required.

5.2.2 The 5‐HT2B Receptor The 5‐HT2B receptor (previously known as the 5‐HT2F receptor) was first identified through its mediation of rat stomach fundus contraction, from where its sequence was subsequently cloned (Foguet et al., 1992). The 5‐HT2B receptor, in comparison with other members of the 5‐HT2 receptor family, has a limited distribution within the brain, with 5‐HT2B receptor‐like immunoreactivity localized on neuronal cells within the cerebellum, lateral septum, dorsal hypothalamus, and medial amygdala (Duxon et al., 1997a). The putative functions of this receptor may include a role in anxiety‐like behaviors, as the marginally selective 5‐HT2B receptor agonist, BW 723C86, is anxiolytic following direct intra‐amygdala injection (Duxon et al., 1997b). In addition, this agonist increased the number of punishments received in the rat Vogel drinking conflict test, an effect blocked by the selective 5‐HT2B receptor antagonist, SB 215505 (Kennett et al., 1998). These studies support a role for the 5‐HT2B receptor in anxiety and this subtype may therefore be a therapeutic target for the treatment of depression and anxiety‐like disorders. Certain phases of sleep may also be regulated by the actions of the 5‐HT2B receptor. For example, administration of the 5‐HT2B receptor antagonist, SB 215505, at the beginning of the light phase, increases the waking period, with a corresponding loss in paradoxical sleep (Kantor et al., 2004). One study has demonstrated that activation of the 5‐HT2B receptor induces hyperphagia (Kennett et al., 1997a), implicating the 5‐HT2B receptor in the regulation of feeding, despite most attention being focused on the role of the 5‐HT2C receptor in feeding (see later). The 5‐HT2B receptor is 5‐HT receptor that may be involved in the generation of migraines. The 5‐HT2B/ 2C agonist, m‐CPP, is known to trigger migraine‐like pain in some individuals, and has the ability to induce

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plasma protein extravasation in the guinea pig; an effect blocked by the selective 5‐HT2B receptor antagonist, LY 202146 (Johnson et al., 2003). Furthermore, this receptor has been shown to be expressed by endothelial cells within meningeal blood vessels, the potential site responsible for migraine generation (Schmuck et al., 1996).

5.2.3 The 5‐HT2C Receptor The 5‐HT2C receptor‐binding site was originally identified in the choroid plexus, and displayed high affinity for [3H]5‐HT (Pazos et al., 1984), resulting in the initial classification within the 5‐HT1 receptor family; high affinity for 5‐HT being a key characteristic to guide classification at the time. The 5‐HT2C receptor sequence was subsequently identified in the rat (Julius et al., 1988). In contrast to the 5‐HT2B receptor, the 5‐HT2C receptor has a widespread distribution throughout the brain. Receptor autoradiographical and immunohistochemical studies have complemented each other, identifying putative sites of receptor expression in the choroid plexus, cortex, amygdala, hippocampus, substantia nigra, caudate nucleus, and cerebellum (e.g., Abramowski et al., 1995). Generally, in situ hybridization has colocalized 5‐HT2C receptor transcripts with the binding sites, suggesting that the receptor is postsynaptic, with the exception of a potentially presynaptic receptor localization in the medial habenula. Activation of the 5‐HT2C receptor is thought to induce membrane depolarization, and may mediate some of the excitatory effects of 5‐HT, for instance in piriform cortical pyramidal neurons (Sheldon and Aghajanian, 1991) and neurons nigral neurons (Rick et al., 1995). More recently, attention has been focused on the role of the 5‐HT2C receptor in regulating the firing activity of dopaminergic neurons, particularly within the VTA and substantia nigra. On the whole, evidence suggests that the 5‐HT2C receptor has a tonic inhibitory control on the firing activity of mesolimbic and mesostriatal dopaminergic neurons. For instance, acute administration of the 5‐HT2B/2C receptor antagonist, SB 206553, increased the rate of firing of neurons in the VTA and substantia nigra, resulting in elevated dopamine release in the nucleus accumbens and striatum (Di Giovanni et al., 1999; Alex et al., 2005). The role of the receptor to modulate nigral–striatal dopamine neurons is controversial, however, since the selective 5‐HT2C receptor antagonist, SB 242084, while increasing firing in the VTA, did not alter nigral neuron activity, and the 5‐HT2C receptor agonist, Ro 60‐0175, decreased basal firing of dopaminergic neurons in the VTA again the nigral neurons were not affected (Di Matteo et al., 1999). A major functional role of the 5‐HT2C receptor is postulated to be the control of feeding, which has incited interest in this subtype as a therapeutic target for antiobesity drugs. The first indication of such a role arose from observations that 5‐HT2C receptor knockout mice were abnormally overweight (Tecott et al., 1995). In addition, the 5‐HT2B/C agonist, mCPP, can induce hypophagia, which is prevented by the 5‐HT2C receptor antagonist, SB 242084 (Kennett et al., 1997b). The anxiolytic properties of 5‐HT2C receptor antagonists suggest a role for this receptor subtype in anxiety (Kennett et al., 1996, 1997b). A more recent report of a new antidepressant compound, agomelatine, which is an agonist of melatonin receptors MT1 and MT2, but also has 5‐HT2C antagonist activity (Den Boer et al., 2006), suggests that blockade of the 5‐HT2C receptor may also have therapeutic benefit in the treatment of depression. Some studies suggest that RNA editing of the 5‐HT2C receptor may be relevant in schizophrenic, depressed, and suicidal patients. This process is known to produce three amino acid alterations within the intracellular regions of the receptor, which may affect the efficiency of the receptor couples with G‐proteins (Burns et al., 1997), and may also influence pharmacology (Niswender et al., 2001). This latter study, however, was unable to detect changes in RNA editing in schizophrenic or depressed patients, though they did identify increased editing at one site in suicide victims. Interestingly, RNA editing appears to be regulated by 5‐HT levels in the brain; perhaps logically depletion attenuates editing, resulting in higher levels of isoforms with enhanced G‐protein‐coupling efficacy (Gurevich et al., 2002). The activity of the 5‐HT2C receptor may, therefore, be altered in diseased states where 5‐HT levels are abnormal, and may lead to further disruption of normal brain functions.

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Similar to the other 5‐HT2 receptor subtypes, the 5‐HT2C receptor may be involved in the regulation of sleep. Antagonists for this receptor promote slow‐wave sleep (Sharpley et al., 1994), though in contrast a more recent study demonstrated that the 5‐HT2C receptor antagonist, SB 242084, increased wakefulness at the expense of deep slow‐wave sleep (Kantor et al., 2005). Clarification of the role of the 5‐HT2C receptor in this respect is clearly required. Due to expression within the hippocampus, the 5‐HT2C receptor has also been postulated to have a role in memory and learning. Of relevance here, a significant impairment in the generation of perforant path‐dentate gyrus LTP, a process considered to be the most likely candidate for the molecular basis of memory, was observed in 5‐HT2C receptor knockout mice (Tecott et al., 1998). In addition, these mice showed impaired spatial learning in the Morris water maze. Finally, activation of the 5‐HT2C receptor may regulate locomotor activity, as the selective agonist, WAY 161503, suppresses measures of spontaneous movement (Mosher et al., 2005).

5.3 The 5‐HT3 Receptor The 5‐HT3 receptor is the only 5‐HT receptor that is a member of the cys–cys loop ligand‐gated ion channel family. The channel is thought to be pentameric (Boess et al., 1995), and may be formed by a combination of up to five different subunits, named 5‐HT3A–E, although at present only the 5‐HT3A and 5‐HT3B subunits have been demonstrated to be incorporated into functional channels. The receptor complex is a nonselective ion channel, permeable to Ca2þ, Naþ, and Kþ ions, that mediates fast synaptic neurotransmission in the brain, via membrane depolarization and is prone to rapid desensitization. Recent attention has focused on the combination of subunits forming the functional channel in native tissue, with current opinion favoring a combination of 5‐HT3A and 5‐HT3B subunits. Expression of only the 5‐HT3A receptor subunit in recombinant systems does not generate a high single‐channel conductance receptor that is evident in some populations of native neuronal receptors, whereas coexpression with the 5‐HT3B receptor modifies the biophysics of the receptor to more resemble some populations of native receptors (Davies et al., 1999; Dubin et al., 1999; > Figure 8-4). In addition to 5‐HT, 5‐HT3 receptor action is modulated, allosterically, by volatile anaesthetics and alcohols (Machu and Harris, 1994; Parker et al., 1996; Suzuki et al., 2002), though the actions of these compounds may depend on the subunit composition of the receptor (Stevens et al., 2005). Within the CNS, the 5‐HT3 receptor is expressed at highest density in the brain stem nuclei, encompassing the chemoreceptor trigger zone; dorsal motor nucleus of the vagus nerve, area postrema, and nucleus tractus solitarius (Pratt et al., 1990). The 5‐HT3 receptor is also found, albeit at much lower levels, in human forebrain regions including the hippocampus, amygdala, and caudate–putamen (Barnes et al., 1989a, b). Interestingly, this latter region in other species (rodents, nonhuman primates) does not display readily detectable expression of 5‐HT3 receptor‐binding sites. A comprehensive understanding of the differential distribution of the individual receptor subunits has not yet been reached, but may be achieved in future studies by the use of selective antibodies recognizing distinct epitopes within the individual subunits. Activation of the 5‐HT3 receptor is believed to modulate the transmission of various neurotransmitters. For example, in the hippocampus, frontal cortex, and hypothalamus, activation of the 5‐HT3 receptor enhances 5‐HT release (e.g., Martin et al., 1992), although the receptor is not thought to be expressed by 5‐HT neurons. In addition, this receptor is thought to have a facilitatory effect on dopamine release. For instance, electrical stimulation of dorsal raphe neurons increases dopamine release in the nucleus accumbens, an effect inhibited by the 5‐HT3 receptor antagonists, ondansetron and zacopride (De Deurwaerdere et al., 1998). Conversely, the 5‐HT3 receptor has an inhibitory effect on acetylcholine release in the cortex (Barnes et al., 1989b) that is likely to be mediated via GABAergic interneurons (Morales and Bloom, 1997; Diez‐Ariza et al., 2002). The primary therapeutic benefit of 5‐HT3 receptor ligands (e.g., ondansetron, granisetron, tropisetron) is their ability to relieve often very severe nausea and vomiting induced by aggressive anticancer chemo‐ and

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. Figure 8-4 The pharmacological and biophysical properties of homomeric and heteromeric 5‐HT3 receptors. (a) Concentration‐dependent activation of currents by 5‐HT recorded from HEK‐293 cells transfected with 5‐HT3A cDNA alone (filled circles) or in combination with 5‐HT3B cDNA (open circles). Data points represent mean current amplitudes recorded from at least four cells normalized to the maximum current amplitude. (b) Concentration‐dependent inhibition by metoclopramide (squares) and tubocurarine (circles) of currents mediated by 5‐HT3A (filled symbols) and heteromeric (open symbols) receptors. (c) Current–voltage relationships for responses evoked by 10 mM 5‐HT, recorded from cells expressing 5‐HT3A (filled circles) and heteromeric (open circles) receptors. Data points represent mean current amplitudes recorded from at least four cells and normalized to the amplitude of the current recorded at 80 mV. Data points for heteromeric receptors were fitted with a linear function. (d) Representative low‐gain d.c. and high‐gain a.c.‐coupled records of an inward current response to 5‐HT (1 mM) recorded at a holding potential of 60 mV from an HEK‐293 cell expressing heteromeric 5‐HT3 receptors. The relationship between membrane current variance and mean current amplitude (1‐s period) was fitted by linear regression for five cells, to yield a single‐channel amplitude (i) of 0.65  0.02 pA and an elementary conductance (g) of 11.7  0.3 pS. (e) Single‐channel recordings from outside‐out patches containing 5‐HT3A (top panel) and heteromeric (lower panel) 5‐HT3 receptors. The conductance (16 pS) of channels mediated by heteromeric receptors was derived from the linear fit to the current–voltage relationship obtained from three excised patches (Data are reproduced with permission from the authors (Davies et al., 1999))

radiotherapy (e.g., Ikeda et al., 2005) and also emesis occurring postoperatively, particularly evident following procedures involving the abdomen (Du Pen et al., 1992). Less well understood is the potential efficacy of 5‐HT3 receptor antagonists to reduce anxiety and other symptoms associated with the forebrain. Despite studies showing that 5‐HT3 receptor agonists may increase anxiety levels in the mouse (Costall et al., 1989), in addition to antagonists having anxiolytic properties, and the 5‐HT3A receptor knockout mouse exhibiting reduced levels of anxiety in the elevated plus maze and

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social interaction tests (Kelley et al., 2003), 5‐HT3 receptor antagonists do not appear to evoke a robust anxiolytic response in patients. The 5‐HT3 receptor may also be involved in cognitive processes, such as learning and memory (Barnes et al., 1990), which may be mediated through the indirect effects of the 5‐HT3 receptor on acetylcholine release. Also of relevance here, activation of the 5‐HT3 receptor has been demonstrated to inhibit long‐term potentiation in the rat hippocampus (Passani et al., 1994); this process being a well‐recognized physiological phenomenon believed to represent memory. Consistent with these findings, 5‐HT3 receptor antagonists have been forwarded for the treatment of various neurological disorders in which memory deficit is concerned, including Alzheimer’s disease, although the clinical responses are not encouraging. Other potential therapeutic roles of 5‐HT3 receptor antagonists include the relief of alcohol addiction (Dawes et al., 2005), the treatment of tardive dyskinesia (Sirota et al., 2000), and the treatment of pain. The role of the 5‐HT3 receptor in this latter process is currently ambiguous due to conflicting reports of its activation being pro‐ or antinociceptive. For example, the selective 5‐HT3 receptor antagonist, ondansetron, inhibits the formalin‐induced response in dorsal horn neurons (Green et al., 2000), while use of the 5‐HT3 receptor agonist 2‐methyl 5‐HT reduces the behavioral response to formalin (Sasaki et al., 2001). The 5‐HT3 receptor antagonists have also been found useful in the treatment of fibromyalgia (e.g., Fa¨rber et al., 2000), a disorder characterized by chronic, widespread pain. In addition, 5‐HT3 receptor antagonists, granisetron, tropisetron, and ondansetron, have been proposed to be beneficial in treating chronic fatigue syndrome (Spa¨th et al., 2000; The et al., 2003).

5.4 The 5‐HT4 Receptor Although only one 5‐HT4 receptor gene has been identified, the arising mRNA can be alternatively spliced within the corresponding C‐terminal region to produce nine isoforms, 5‐HT4A–H and 5‐HT4HB, although it is possible that more may become apparent. Most isoform transcripts studied appear to be expressed in the brain, with the exception of 5‐HT4D, which currently has been located only in the gut. The isoforms do not appear to differ pharmacologically, though they may vary in G‐protein‐coupling efficiency (Mialet et al., 2000), perhaps not surprising given the putative role of the C terminus of GPCRs to interact with G‐protein subunits. Various studies have investigated the location of the 5‐HT4 receptor within the brain, the binding sites and mRNA colocalize, indicating a probable postsynaptic location. Receptor autoradiography using human tissue has identified 5‐HT4 receptors with highest levels in the basal ganglia, including the substantia nigra, globus pallidus, caudate nucleus, putamen, nucleus accumbens, hippocampus (CA1 and subiculum), and cortex (Varnas et al., 2003). The 5‐HT4 receptor is positively coupled to adenylate cyclase, and enhances neuronal excitability (Gerald et al., 1995). Consistent with this role, the 5‐HT4 receptor increases the release of acetylcholine in the rat frontal cortex, which is activated using the selective 5‐HT4 receptor agonists, BIMU1 and BIMU8 (Consolo et al., 1994). Although antagonists block this response, they have no effect on acetylcholine release alone, suggesting that the 5‐HT4 receptor has little tonic control of basal cholinergic transmission. The 5‐HT4 receptor has also been shown to have a facilitatory effect on striatal dopaminergic transmission (Steward et al., 1996). Likewise, in 1993, Benloucif et al. (Benloucif et al., 1993) demonstrated that the nonselective 5‐HT4 receptor agonist, 5‐MT, could increase dopamine release in the striatum. More recently, the morphine‐induced increase in striatal dopamine release was inhibited by two selective 5‐HT4 receptor antagonists, GR 125487 and SB 204070. These compounds exerted no effect in the absence of neuronal excitation, or following amphetamine‐induced dopamine release, suggesting that only this receptor subtype has a modulatory influence during dopaminergic neuron activity (Porras et al., 2002). Interestingly, the 5‐HT4 receptor also facilitates nigral dopamine release (Thorre´ et al., 1998). 5‐HT release in the hippocampus (Ge and Barnes, 1996) also appears to be enhanced by the 5‐HT4 receptor, with the receptor being tonically active in freely moving rats. It has been suggested that this receptor can regulate the firing activity of serotonergic neurons originating in the dorsal raphe nucleus (Lucas et al., 2005). Such manipulation of the central 5‐HT system may forward this receptor as a target for antidepressant therapy.

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The 5‐HT4 receptor is believed to have a major role in mediating the neuronal basis of learning and memory. Ligands acting on this receptor subtype, therefore, have been studied to identify any cognitive‐ enhancing properties that may prove beneficial in the treatment of conditions such as Alzheimer’s disease. Moreover, there has been a report of reduced 5‐HT4 receptor density in the postmortem brains of Alzheimer’s disease sufferers (Reynolds et al., 1995), implying that the receptor may be involved in the pathogenesis of this disorder. Many studies have shown that 5‐HT4 receptor activation enhances performance in numerous behavioral paradigms of cognitive function. For example, the 5‐HT4 receptor agonist, RS 17017, improved primate performance in the delayed matching task (Terry et al., 1998), whereas an alternative 5‐HT4 receptor agonist, RS 67333, enhanced learning in the Morris water maze (Lelong et al., 2001). The partial 5‐HT4 receptor agonist, SL65.0155, also facilitated retention during an object recognition task, which was antagonized by the 5‐HT4 receptor antagonist, SDZ 205557 (Moser et al., 2002). The same report described a synergistic effect on memory enhancement when combining SL65.0155 and the cholinesterase inhibitor, rivastigmine, further implying that use of 5‐HT4 receptor agonists may have beneficial effects in treating Alzheimer’s disease. These positive effects of 5‐HT4 receptor activation may be mediated by its regulation of acetylcholine release in the cortex. An alternative possibility is the interaction of this receptor with amyloid precursor protein (APP) metabolism. 5‐HT4 receptor agonists appear to increase the secretion of sAPPa (> Figure 8-5), a neuroprotective peptide which counteracts the cellular toxicity generated by the overactivity of glutamatergic transmission, promoting neuronal growth. Indeed, some studies suggest that this polypeptide can enhance memory functions in behavioral paradigms. 5‐HT4 receptor activation may, therefore, improve cognitive functions by facilitating the release of this neuroprotective peptide (Lezoualc’h and Robert, 2003). In addition, the 5‐HT4 receptor may enhance cognitive performance through potentiating hippocampal LTP, the cellular mechanism proposed to underlie memory. Indeed, it has been demonstrated that activation of the 5‐HT4 receptor induces depolarization of pyramidal cells within the CA1 field (Chapin et al., 2002). The 5‐HT4 receptor may also have a role in the generation of anxiety. 5‐HT4 receptor antagonists, for example, have been shown to exhibit anxiolytic properties (Kennett et al., 1997c), whereas the 5‐HT4 receptor knockout mouse exhibited abnormal responses to stress, whereby stress‐induced hypophagia was attenuated in the knockout mouse compared with the wild‐type strain (Compan et al., 2004). Further investigation of 5‐HT4 receptor antagonists may, therefore, identify new compounds to treat anxiety.

5.5 The 5‐ht5 Receptors The 5‐ht5 subfamily contains the 5‐ht5A and 5‐ht5B receptors and despite nearly 15 years since their discovery, they remain the least well understood 5‐HT receptor subtypes with no conclusive evidence as to how they elicit second‐messenger responses despite their structural classification as GPCRs. Physiological roles for these receptor subtypes have not been identified, hence they retain lower‐case appellation to emphasize their current status as gene products, contrasting with the upper‐case notation of a receptor with known cellular functions. Of the two subtypes, the 5‐ht5A receptor has been the focus of most investigations. Currently, opinion favors negative coupling of the 5‐ht5A receptor to adenylate cyclase within recombinant cell systems (Francken et al., 1998; Hurley et al., 1998), though other reports have detected no such response (Grailhe et al., 2001). It has also been suggested that the 5‐ht5A receptor may induce intracellular Ca2þ mobilization (Noda et al., 2003) or couple to an inwardly rectifying potassium channel (Grailhe et al., 2001). In 2000, Oliver et al. (Oliver et al., 2000) conducted the first extensive study of 5‐ht5A receptor protein expression in the rat brain using immunohistochemistry. Immunoreactivity appeared to be associated with neurons, and was most strongly identified in the hypothalamus, raphe nuclei, locus coeruleus, horizontal nucleus of the diagonal band, and amygdala, with moderate staining in many regions of the cortex (particularly entorhinal cortex), the hippocampus, lateral habenula, substantia nigra, VTA, pons, and cerebellum. In situ hybridization using human brain tissue has demonstrated 5‐ht5A receptor transcripts in the cortex, hippocampus, amygdala, and cerebellum (Pasqualetti et al., 1998).

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. Figure 8-5 Activation of the h5‐HT4(g) receptor increases the release of nonamyloidogenic sAPPa. (a and b) Representative immunoblots showing the effects of increasing concentrations of 5‐HT (a) and prucalopride (b) on the cellular release of sAPPa in CHO cells stably coexpressing the h5‐HT4(g) receptor and human APP695. After incubating the cells with the indicated concentrations of ligands for 30 min, secreted sAPPa was measured by Western blot using the monoclonal antibody 6E10. A 110‐kDa molecular weight marker is indicated at the right. (c) Representative immunoblot showing the blocking effect of the selective 5‐HT4 antagonist, GR 133808, on 5‐HT and pruclapride‐induced sAPPa release. CHO cells were preincubated with 1 mM concentration of GR113808 10 min before treatment with 5‐HT4 ligands. After an additional 30‐min period, sAPPa was detected in the culture medium. Immunoblots were performed as in (a) and (b). CT, untreated control cells. Experiments were repeated at least three times with similar results (Data are reproduced with permission from the authors (Lezoualc’h and Robert, 2003))

Although no definitive role for the 5‐ht5A receptor has yet been elucidated, hindered greatly by the lack of an available selective ligand although this position appears to be changing (Corbett et al., 2005; Thomas, 2006), a few studies have suggested putative functions. The 5‐ht5A receptor knockout mouse, for example, exhibited increased levels of exploratory behavior in response to a novel environment as well as an attenuated response to the nonselective 5‐HT receptor ligand, LSD, suggesting that the 5‐ht5A receptor may mediate some of the behavioral effects elicited by this drug of abuse (Grailhe et al., 1999). The 5‐ht5A receptor may also be involved in the regulation of circadian rhythm in rats, although overlapping pharmacology with the 5‐HT7 receptor complicates interpretation (Sprouse et al., 2004). The development of compounds with selectivity for the 5‐ht5A receptor will greatly facilitate the elucidation of a physiological role for this receptor subtype. One such compound reported recently, SB 699551‐A (Corbett et al., 2005; Thomas, 2006), exhibits a 30‐fold selectivity for the human 5‐ht5A receptor over other 5‐HT receptor subtypes and other neurotransmitter receptors, aside from the serotonin transporter, for which it has only a

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tenfold selectivity (Thomas et al., 2006). Unfortunately, it appears that unlike the human and guinea pig receptors, this compound has a low affinity for the mouse and rat 5‐ht5A receptors (pKi ¼ 6.3), which limits the value of this compound in elucidating 5‐ht5A receptor function through common rodent paradigms. The 5‐ht5B receptor has attracted even less attention than the 5‐ht5A receptor, no doubt due to the discovery that the human 5‐ht5B gene sequence contains stop codons within its open reading frame, and therefore is not expected to encode a functional protein (Grailhe et al., 2001). The rat and mouse 5‐ht5B receptors, however, appear to be expressed and may be functional, though no evidence has been presented to support this latter notion. Identification of 5‐ht5B mRNA in the rat brain demonstrated expression in the hippocampus, habenula, entorhinal and piriform cortices, and the olfactory bulb (Matthes et al., 1993).

5.6 The 5‐HT6 Receptor The sequence for the 5‐HT6 receptor was first identified in 1993 (Monsma et al., 1993; Ruat et al., 1993) yet in terms of functional significance, this receptor remains relatively poorly understood. Investigations into the location of the receptor have identified receptor transcripts in the striatum, nucleus accumbens, hippocampus, and olfactory tubercles of rat and guinea pig brain tissue (Ruat et al., 1993; Ward et al., 1995). Correspondingly, the 5‐HT6 receptor protein has been identified using selective antibodies in the cortex (frontal, entorhinal, and piriform), nucleus accumbens, cerebellum, caudate putamen, substantia nigra, hippocampus (in particular, the dentate gyrus and CA1), and olfactory tubercles (Gerard et al., 1997; Hamon et al., 1999). Higher resolution studies using electron microscopy have identified the ultrastructural distribution of the protein, which is expressed postsynaptically by dendrites in the hippocampus and striatum (Hamon et al., 1999). It should be noted that while rat and human tissue readily express the 5‐HT6 receptor, the mouse brain appears to express very low levels of the receptor, thus most studies attempting to define functions for the 5‐HT6 receptor have used rat models (Hirst et al., 2003). Although the 5‐HT6 receptor is found predominantly in the CNS, it has also been detected nonneuronally within the thymus, spleen, and lymphocytes (Stefulj et al., 2000). The 5‐HT6 receptor has been demonstrated to be positively coupled to adenylate cyclase in both recombinant systems and in pig striatal tissue (Monsma et al., 1993; Schoeffter and Waeber, 1994). The receptor is thought to modulate the activity of various neurotransmitter systems, including acetylcholine, dopamine, noradrenaline, and glutamate, although some of the evidence currently appears to be conflicting. Inhibition of the 5‐HT6 receptor appears to facilitate cholinergic transmission, with evidence arising from studies using antisense oligonucleotides to knock down receptor expression, which produced a behavioral syndrome consisting of yawning, chewing, and stretching (Bourson et al., 1995). These behavioral responses were blocked with the muscarinic acetylcholine receptor antagonist, atropine. Reassuringly, a similar behavioral syndrome was induced by the 5‐HT6 receptor antagonist, Ro 04‐6790, which was also blocked by the muscarinic acetylcholine receptor antagonists (Bentley et al., 1999), implicating the central acetylcholine system in the response (Bentley et al., 1999). 5‐HT6 receptor blockade may also facilitate dopaminergic transmission. Despite some studies finding that 5‐HT6 receptor antagonists have no effect on dopaminergic transmission (Dawson et al., 2001), others have found that the administration of a 5‐HT6 receptor antagonist appears to regulate an enhanced dopaminergic system, as occurs following amphetamine administration (Pullagurla et al., 2004). In addition, increased levels of extracellular dopamine occur in the rat mPFC following administration of the 5‐HT6 receptor antagonist, SB 271046 (Lacroix et al., 2004). The 5‐HT6 receptor also regulates glutamatergic and GABAergic neurotransmission. Thus extracellular levels of glutamate in the frontal cortex and dorsal hippocampus increase following SB 271046 administration (Dawson et al., 2001), while the 5‐HT6 receptor agonist, WAY 446, elevates GABA levels in the cortex and hippocampus (Schechter et al., 2004). These findings are consistent with the identified cellular expression of the receptor (Hamon et al., 1999). The putative ability of the 5‐HT6 receptor to modulate acetylcholine release has attracted attention due to the role of this system in cognitive processes (see Mitchell and Neumaier, 2005, for a review). Generally, it is believed that 5‐HT6 receptor antagonists have a positive effect on cognition. For example, it has been

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shown that antagonists can relieve the amnesia induced by an anticholinergic drug (e.g., Woolley et al., 2003), though other reports have not been able to identify this response (Lindner et al., 2003). Additional studies support the positive effects of 5‐HT6 receptor antagonists on cognitive processes. For example, the 5‐HT6 receptor antagonist, Ro 04‐6790, enhanced the retention of spatial learning in the Morris water maze (Woolley et al., 2001) and consolidation in the novel‐object discrimination task (King et al., 2004), while the selective antagonists, SB 357134 and SB 399885, improved memory consolidation in an autoshaping learning task (Perez‐Garcia and Meneses, 2005). These studies suggest that 5‐HT6 receptor antagonists may have therapeutic value in reversing cognitive deficits and the results of clinical studies are eagerly awaited. A role for the 5‐HT6 receptor in the treatment of schizophrenia has also been suggested, largely due to the observation that some atypical antipsychotic compounds, including olanzapine and clozapine, have appreciable affinity for the 5‐HT6 receptor. Indeed, chronic administration of clozapine has been shown to reduce 5‐HT6 receptor mRNA levels in the rat hippocampus (Frederick and Meador‐Woodruff, 1999; > Figure 8-6). Action of these drugs at this subtype may, therefore, be responsible for some of the

. Figure 8-6 Effect of clozapine and haloperidol on 5‐HT6 mRNA expression in hippocampus. There was a significant effect of treatment. Post hoc analysis revealed that the treatment effect was entirely due to clozapine (clozapine vs. control, p < 0.01; haloperidol vs. control, p > 0.15) (Data are reproduced with permission from the authors (Frederick and Meador‐Woodruff, 1999))

therapeutic effects, though it may be argued that such interaction may mediate some of the adverse effects of the drug. In further support of a role for the 5‐HT6 receptor in the pathogenesis of schizophrenia, levels of mRNA appear to be reduced in the hippocampus of schizophrenic patients, though the density of 5‐HT6 receptor‐binding sites appears not to vary (East et al., 2002a, b). In addition, a study of Ro 04‐6790 on prepulse inhibition (PPI) in rats, a measure known to be attenuated in schizophrenia, showed no effect, hence failing to offer support that this receptor is involved in schizophrenia (Leng et al., 2003). Many association studies have been performed, analyzing putative links between polymorphisms in the 5‐HT6 receptor gene and schizophrenia. No clear association has been identified as the results have been conflicting (e.g., Shinkai et al., 1999; Tsai et al., 1999), which is often the case in such studies. The 5‐HT6 receptor has also been associated with the pathogenesis or treatment of anxiety and depression. For instance, treatment with antisense oligonucleotides resulted in a loss of 5‐HT6 receptor expression in the nucleus accumbens, in addition to enhanced anxiety levels evident in the elevated plus maze and social interaction tests (Hamon et al., 1999), which may be indicative of 5‐HT6 receptor

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activation having an antidepressant‐like effect. Furthermore, 5‐HT6 receptor expression may be regulated by stress hormones like corticosteroids. Thus prevention of corticosteroid release increases 5‐HT6 receptor mRNA levels in the hippocampus (Yau et al., 1997), suggesting that 5‐HT6 receptor activity may fall in response to stress, and hence might contribute to the generation of depression. Other potential therapeutic benefits of 5‐HT6 receptor ligands may include the control of seizures, as the receptor antagonist, SB 271046, displays anticonvulsant properties, though this effect was small in comparison with the efficacy of better recognized antiepileptic drugs (Routledge et al., 2000). Additionally, the 5‐HT6 receptor may be a target for antiobesity drugs, as both antisense oligonucleotide knock down of the receptor and the 5‐HT6 receptor antagonist, Ro 04‐6790, reduced rat body weight (Woolley et al., 2001).

5.7 The 5‐HT7 Receptor Unlike the other, more recently identified 5‐HT receptors, the 5‐HT7 receptor has been the subject of numerous investigations providing evidence as to its function. Although there is only one 5‐HT7 receptor gene, four distinct isoforms are generated through alternative splicing generating variations within the C terminus of the polypeptide sequence (Heidmann et al., 1997). Hence, it is possible that the different isoforms have differing abilities to couple to second‐messenger systems, though one report suggests that the three human isoforms have indistinguishable pharmacological and coupling characteristics (Krobert et al., 2001). In general, the distribution of 5‐HT7 receptor transcripts and receptor‐binding sites are similar, indicating a postsynaptic expression. A receptor autoradiographical study with human brain tissue demonstrated the distribution of 5‐HT7 receptor‐binding sites using the [3H] derivative of the selective 5‐HT7 receptor antagonist, SB 269970 (Varnas et al., 2004). 5‐HT7 receptor expression appears to be the highest in the thalamus and dentate gyrus of the hippocampus, with lower expression in the substantia nigra, VTA, dorsal raphe nucleus, Ammon’s horn of the hippocampus, cingulate cortex, amygdala, and hypothalamus. Low levels of expression have also been found in the cortex, subiculum, and parahippocampal gyrus. In the human brain, mRNA levels of 5‐HT7(a) and 5‐HT7(b) are comparable in the caudate and hippocampus, while the 5‐HT7(a) isoform is the predominant species in rat. In contrast, 5‐HT7(c) (rat) and 5‐HT7(d) (human) are expressed at relatively low levels (Heidmann et al., 1998). The 5‐HT7 receptor has been demonstrated to be positively coupled to adenylate cyclase (Bard et al., 1993), and modulates neuronal activity in various brain regions. For example in the hippocampus, activation of the 5‐HT7 receptor is believed to increase neuronal activity. For instance, the nonselective 5‐HT7 receptor agonist, 5‐CT, in the presence of WAY 100635 to block action at 5‐HT1A receptors, mediates an increase in the amplitude of the population spike (Tokarski et al., 2003). In addition, recoding from individual pyramidal neurons, 5‐HT7 receptor activation inhibits the slow AHP (sAHP) in both CA1 and CA3 cells (Bacon and Beck, 2000; Tokarski et al., 2003). A 5‐HT7 receptor‐mediated inhibition of the sAHP has also been identified in the thalamus (Goaillard and Vincent, 2002), where the 5‐HT7 receptor is also thought to increase neuronal excitability through modification of Ih, the hyperpolarization‐activated nonselective cation current (Chapin and Andrade, 2001). Within the DRN, the 5‐HT7 receptor may have a negative influence on the firing of 5‐HT neurons (Harsing et al., 2004). These authors suggested that this effect was due to the expression of 5‐HT7 heteroreceptors on the terminals of glutamatergic cortico‐raphe neurons. Alternatively, the 5‐HT7 receptor may facilitate raphe 5‐HT neuron activity via GABAergic interneurons. Roberts et al. (2004) demonstrated that the 5‐HT efflux in the guinea pig DRN was inhibited by the selective 5‐HT7 receptor antagonist, SB 269970‐A; the action was blocked by the GABAA receptor antagonist, bicuculline. They postulated that activation of the 5‐HT7 receptor induces a reduction in GABA release, hence facilitating serotonergic neuron activity. The 5‐HT7 receptor, probably along with the 5‐HT1A receptor, mediates the hypothermia induced by the nonselective agonists, 5‐CT and 8‐OH‐DPAT, since the response can be antagonized by the selective 5‐HT7 receptor antagonist, SB 269970 (Hagan et al., 2000). The 5‐HT7 receptor also induces phase shifts in the circadian rhythm of neurons within the SCN, where the receptor is expressed (Duncan et al., 1999). These 8‐OH‐DPAT‐induced phase shifts are antagonized by SB 269970 (Duncan et al., 2004). The 5‐HT7

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receptor may also regulate various neuroendocrine functions, such as the release of vasopressin and oxytocin (Jorgensen et al., 2003). In addition to circadian rhythm, it is also likely that the 5‐HT7 receptor regulates certain sleep phases, thus the 5‐HT7 receptor knockout mouse displays reduced levels of rapid eye movement (REM) sleep (Hedlund et al., 2005; > Figure 8-7), consistent with a reduction in REM sleep evident following administration of a selective 5‐HT7 receptor antagonist (Thomas et al., 2003). A recent study has investigated this action further by directly administering the antagonist SB 269970 to the DRN, which reduced REM sleep as a whole and also the number of REM periods (Monti and Jantos, 2006). The ability of the GABAA receptor

. Figure 8-7 Time courses of wakefulness, slow‐wave sleep, and rapid eye movement sleep over a 24‐h period. Both 5‐HT7þ/þ (open square) and 5‐HT7/ (filled square) mice showed a normal circadian sleep pattern. During the light period, 5‐HT7/ mice spent less time in rapid eye movement sleep. There was no difference between the genotypes in wakefulness or slow‐wave sleep. Values are expressed as mean  SEM. n ¼ 10 animals/group, *p < .05, two‐way repeated measures analysis of variance followed by a Bonferroni test (Data are reproduced with permission from the authors (Hedlund et al., 2005))

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agonist, muscimol, to prevent this response led the authors to suggest that the effects were mediated via GABAergic transmission within the DRN. The 5‐HT7 receptor is a putative therapeutic target for various other CNS disorders, including depression. Studies with the 5‐HT7 receptor knockout mouse have demonstrated a reduced immobility in forced‐swim tests and tail suspension tests, similar to behaviors exhibited by wild‐type mice following treatment with antidepressant agents (Guscott et al., 2005; Hedlund et al., 2005). Also of potential relevance, chronic treatment with the SSRI antidepressant compound, fluoxetine, and the tricyclic antidepressant drug, imipramine, reduces 5‐HT7 receptor‐binding sites in the hypothalamus (Sleight et al., 1995; Mullins et al., 1999). Additionally, treatment with citalopram and imipramine can reduce the excitatory effect of 5‐HT7 receptor activation on hippocampal bursting frequency (Tokarski et al., 2005). These studies suggest that antidepressants may exert beneficial effects through a potential downregulation of 5‐HT7 receptors. The evidence for a potential role for the 5‐HT7 receptor in schizophrenia is much less persuasive, being solely based on the affinity of some antipsychotic compounds having a high affinity for the 5‐HT7 receptor. Evidence against such a role has been provided by East et al. (1999), who could not find any difference in hippocampal 5‐HT7 receptor mRNA levels between control and schizophrenic tissue. Conversely, putative 5‐HT7 receptor involvement in epilepsy appears to be more promising. A rat model of absence epilepsy, WAG/Rij, displayed reduced epileptic activity following administration of the selective 5‐HT7 receptor antagonist, SB 258719 (Graf et al., 2004). The high level of 5‐HT7 receptor expression in the hippocampus has led to the suggestion that this receptor subtype may have a role in learning and memory. Interestingly, the 5‐HT7 receptor knockout mouse displays an impaired response to contextual fear conditioning (Roberts et al., 2004). The same report also demonstrated a reduced occurrence of LTP in the CA1 field of 5‐HT7 receptor knockout mice. Furthermore, 5‐HT7 receptor mRNA expression is increased in the hippocampus and PFC of rodents having undertaken autoshaping–training, a model of learning (Perez‐Garcia et al., 2006). Further investigation of the role of the 5‐HT7 receptor in cognitive processes may lead to the development of new clinically effective compounds in the treatment of neurological disorders.

6

The 5‐HT Transporter (SERT)

The 5‐HT transporter (SERTor 5‐HTT) is a member of the Naþ/Cl‐dependent biogenic amine transporter family, which includes the dopamine (DAT) and noradrenaline (NET) transporters (see Masson et al., 1999, for review). SERT plays a vital role within the 5‐HT system, limiting 5‐HT neurotransmission by removing the neurotransmitter through transport across the presynaptic membrane (Rudnick and Clark, 1993). Following the sequencing of rat SERT (Blakely et al., 1991), subsequent studies have suggested that SERT may exist as an oligomer in vivo. Ramamoorthy et al. (1993), for example, demonstrated that the predicted molecular weight of the transporter is
70 kDa, which contrasted with the 300 kDa estimated from the purified human placental protein. Furthermore, the transporter forms homooligomers in vitro (Kilic and Rudnick, 2000). Within the brain, SERT is located presynaptically on 5‐HT neurons, and displays central distribution closely matching the regions receiving 5‐HT neuron innervation. In situ hybridization studies in the rat demonstrated that SERT transcript was present in 5‐HT neuron cell bodies of most of the raphe nuclei in the hindbrain (Fujita et al., 1993). Correspondingly, human postmortem brain tissue autoradiography using the [3H] derivative of the tricyclic antidepressant, imipramine exhibited highest levels of binding in the raphe nuclei and midline thalamic nuclei, with weaker signals in the substantia nigra, locus coeruleus, nucleus interpeduncularis, nucleus nervi hypoglossi, nucleus nervi facialis, mammillary bodies and other regions of the hypothalamus, and low levels in the cortex, hippocampus, and amygdala (Cortes et al., 1988). Use of the SSRI, [3H]paroxetine yielded binding at highest levels in the substantia nigra, hypothalamus, and hippocampus, with lower levels in the basal ganglia and thalamus (Laruelle et al., 1988). Immunohistochemical investigation of SERT protein location in the rat brain has identified expression in the raphe nuclei, B9 and throughout the forebrain, including the hypothalamus, hippocampus, substantia nigra,

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amygdala, and cortex (Qian et al., 1995), in addition to the spinal cord (Sur et al., 1996). The location of SERT in these studies corresponded with 5‐HT immunoreactivity. The subcellular location of SERT appears to be dendritic and axonal, in addition to somal in the raphe nuclei (Qian et al., 1995), though some studies have been unable to demonstrate perikaryal labeling of neurons within this region (Yamamoto et al., 1998). Furthermore, electron microscopy has confirmed the predicted presynaptic location for the transporter in the rat cortex (Yamamoto et al., 1998). Despite exhibiting a predominant neuronal localization, it has been postulated that SERT may be expressed by glial cells (Lawrence et al., 1995a, b), though other reports have been unable to identify such expression (Fujita et al., 1993; Qian et al., 1995). A recent study has suggested that more than one form of SERT protein is present in vivo. Shigematsu et al. (2006) conducted immunohistochemical studies on the mouse brain with two selective antibodies, one raised against an epitope within the C terminus, the other against part of the N terminus. They observed that immunoreactivity with the N‐terminal antibody was absent in some regions, notably within the CA3 field of the hippocampus, where the C‐terminal antibody was observed to indicate SERT expression. This implies that SERT may contain variable N‐terminal domains, potentially through alternative splicing of exon 1. Interestingly, despite SERT being found predominantly, if not totally, on serotonergic neurons in the adult brain, expression of the transporter has been found to occur transiently in glutamatergic thalamocortical afferents in the developing mouse brain (Lebrand et al., 1996; Bruning and Liangos, 1997). These neurons are also immunopositive for 5‐HT, and since they are unable to synthesize this neurotransmitter, it has been suggested SERT sequesters 5‐HT, enabling the afferents to mediate serotonergic transmission during brain development. Expression of SERT within these neurons is believed to be maintained for several postnatal weeks (D’Amato et al., 1987). The actions of antidepressant drugs, in particular, the SSRIs, have led to extensive research attempting to elucidate the physiological roles of SERT in the brain. It seems indisputable that the transporter is involved in depression, though its precise mechanism of action is presently unclear. Many studies have investigated the genetic variation that occurs upstream of the SERT‐coding sequence, within the region known as 5‐HTT gene‐linked polymorphic region (5‐HTTLPR), to link the transporter with the occurrence of depression. This sequence consists of a series of repeated units, and acts as a promoter region to regulate levels of SERT expression (Lesch et al., 1996; Greenberg et al., 1999). A common polymorphism within this region is a 44‐base pair deletion, which gives rise to a short form (S), and two variations of a long form (LG and LA). The presence of the short‐form allele reduces SERT expression and activity in vitro, in contrast to cells homozygous for the long allele (Lesch et al., 1996), while a heterozygous genotype is associated with reduced SERT mRNA levels compared with a long allele genotype in the human brain (Little et al., 1998). More recently, it has been demonstrated that LG also results in levels of SERT expression comparable with the short‐form variant (Hu et al., 2006). It has been suggested that individuals carrying at least one short‐ form allele are predisposed to depressive episodes. Hariri et al. (2002) demonstrated increased neuronal activation in the human amygdala in response to fear when the short‐form allele was present. Furthermore, Caspi et al. (2003) monitored the occurrence of stressful life events in young adults and any subsequent depressive episodes, and found that the presence of the short allele was associated with a much greater incidence of depression when stressful life events were experienced. In conjunction, reduced levels of SERT binding in the brains of living depressed patients have been detected in the brain stem (Malison et al., 1998), amygdala, and midbrain (Parsey et al., 2006b), when compared with control subjects using SPECT and PET. Perhaps importantly, the reduction in SERT levels was greater in drug‐naı¨ve patients. In contrast, it does not appear that the short allele is associated with reduced SERT levels in the adult brain (Parsey et al., 2006a), though it cannot be ruled out that this polymorphism has an effect on SERT levels at earlier stages of life. Depression in monkeys, modeled through maternal separation, has also been linked with a reduction in SERT‐binding potential within the raphe nuclei and numerous forebrain regions assessed using PET analysis, suggesting that stress in early stages of life may lead to alterations in adult levels of SERT (Ichise et al., 2006). Studies of SERT function through knockout mouse strategies have also detected behavioral abnormalities that may be related to depression and anxiety. In 2003, Lira et al. (Lira et al., 2003) were unable to detect any differences in anxiety levels between SERT knockout and wild‐type strains of mice using the elevated plus maze and open‐field tests, but did identify abnormalities in the behavioral responses during

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paradigms designed to detect the antidepressant actions of compounds; increased immobility in the forced‐ swim test, higher rates of escape failure in shock avoidance, and longer periods of novelty‐suppressed feeding all imply a depressive‐like state. These differences may be explained by a 50% reduction in DRN neuron number and a fourfold reduction in neuron firing rate in the knockout mice. A further study also suggests that SERT knockout strains exhibit depressive or despair‐like states determined by an increase in the immobility in the tail suspension test (Zhao et al., 2006). These authors also reported a potential increase in anxiety levels. Furthermore, Adamec et al. (2006) postulate that loss of SERT may render an individual vulnerable to posttraumatic stress disorder, as SERT knockout mice displayed increased anxiety in response to predator odor exposure. Aside from a potential role in depression, SERT may be involved in other psychological disorders, including suicidal states. However, various reports have detected increased, decreased, and unchanged levels of SERT in the brains of suicide victims using autoradiographical approaches, resulting in no current consensus as to whether SERT has a role in the pathogenesis of suicidal behavior (see Purselle and Nemeroff, 2003, for an extensive review). Likewise, attempts to link the long and short alleles of the 5‐ HTTLPR locus with suicide have been similarly contradictory (Purselle and Nemeroff, 2003), thus further study is required. SERT may also have a role in the pathogenesis or treatment of schizophrenia. It has been observed that administration of SSRIs to schizophrenic patients results in the improvement of negative symptoms (Silver et al., 2000), an effect apparently mediated independently of antidepressant mechanisms, as use of an antidepressant with no effect on serotonin reuptake (maprotiline) was ineffective (Silver and Shmugliakov, 1998). Investigations of SERT‐binding sites in postmortem tissue, however, have been unable to generate a consensus as to whether levels of the transporter are aberrant in schizophrenic patients. Within the frontal cortex, for example, one study has reported no differences in the density of 5‐HT uptake sites (Gurevich and Joyce, 1997). Other potential sites of SERT‐binding changes include the caudate, hippocampus, nucleus accumbens, and putamen, though further studies are necessary before drawing of definitive conclusions. One report employed the use of PET to determine changes in SERT levels of living schizophrenic patients, and demonstrated no differences in any region assessed (midbrain, thalamus, caudate, putamen, striatum, amygdala, entorhinal cortex, and hippocampus), though the authors suggest that any changes in cortical SERT‐binding levels may be masked due to low expression levels in this region (Frankle et al., 2005). Another psychological disorder involving SERT activity may be obsessive compulsive disorder (OCD), as genetic studies have identified a potential link between OCD and the long‐form 5‐HTTLPR allele, LA (Hu et al., 2006). Recently, SERT has become an interesting potential therapeutic target in epilepsy research, as the use of two SSRIs, fluoxetine and citalopram, were shown to reduce seizures in nonsymptomatic epilepsy sufferers, completely abolishing seizures in approximately one‐third of the subjects (Albano et al., 2006). These results have tremendous potential, and study of putative SERT abnormalities in epileptic brains may identify a role for the transporter in the pathogenesis of seizures. In contrast with its role in treating various neurological disorders, SERT is also a target for various drugs of abuse, including MDMA (ecstasy) and cocaine. MDMA, for example, targets SERT, blocking 5‐HT reuptake and enhancing 5‐HT release (Pletscher et al., 1963; Rudnick and Wall, 1992). While cocaine is predominantly considered as acting on DAT, it is thought that SERT interaction contributes to the reward effects of cocaine (Rocha et al., 1998); hence, pharmacological manipulation of SERT activity may be exploited to prevent drug abuse through reward pathway inhibition.

7

Conclusions

The serotonergic system within the brain has been the subject of extensive research over the past five decades, resulting in a wealth of information regarding its anatomy, the receptors that mediate 5‐HT transmission, and the regulation of transmission through the 5‐HT reuptake transporter. Despite this intensity of activity, there remain numerous gaps in our understanding such as the possible functions of a number of the receptors (e.g., 5‐HT1D, 5‐ht1E, 5‐HT1F, 5‐ht5A, and 5‐ht5B receptors), the physiological significance of RNA editing of the 5‐HT2C receptor, alternative splicing of the 5‐HT4 and 5‐HT7 receptors,

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and the putative existence of homo‐ and heteromeric dimers in vivo – all have the potential to generate a vast diversity of responses to 5‐HT in the brain, and may with time, allow further manipulation of the central 5‐HT system for therapeutic benefit. The potential of each 5‐HT receptor subtype to mediate the pathophysiology of disease and psychological disorders has been discussed in this chapter, and it is already clear that the central 5‐HT system provides a number of important (potential) drug targets ranging from those to treat affective disorders such as anxiety and depression, to those relieving the symptoms of psychosis, Alzheimer’s disease, and emesis. It is perhaps not surprising, therefore, that the pharmaceutical industry maintains an intense interest in the actions of 5‐HT, and we await the development of novel therapeutics resulting from this activity.

References Abramowski D, Rigo M, Duc D, Hoyer D, Staufenbiel M. 1995. Localization of the 5‐hydroxytryptamine 2C receptor protein in human and rat brain using specific antisera. Neuropharmacology 34: 1635-1645. Adamec R, Burton P, Blundell J, Murphy DL, Holmes A. 2006. Vulnerability to mild predator stress in serotonin transporter knockout mice. Behav Brain Res 170: 126-140. Adham N, Kao HT, Schechter LE, Bard J, Olsen M, et al. 1993. Cloning of another human serotonin receptor (5‐HT1F): A 5th 5‐HT1 receptor subtype coupled to the inhibition of adenylate cyclase. PNAS 90: 408-412. Aghajanian GK. 1990. Use of Brain Slices in the Study of Serotoninergic Pacemaker Neurons of the Brainstem Raphe Nuclei. New York: John Wiley & Sons. Aghajanian GK. 1995. Electrophysiology of serotonin receptor subtypes cpathways. Bloom FR, Kupfer DJ, editors. Psychopharmacology: The Fourth Generation of Progress. Raven; New York: pp. 1451-1459. Aghajanian GK, Haigler HJ. 1974. L‐Tryptophan as a selective histochemical marker for serotonergic neurons in single‐ cell recording studies. Brain Res 81: 364-372. Aghajanian GK, Marek GJ. 1999a. Serotonin and hallucinogens. Neuropsychopharmacology 21: S16-S23. Aghajanian GK, Marek GJ. 1999b. Serotonin–glutamate interactions: A new target for antipsychotic drugs. Neuropsychopharmacology 21: S122-S133. Albano C, Cupello A, Mainardi P, Scarrone S, Favale E. 2006. Successful treatment of epilepsy with serotonin reuptake inhibitors: Proposed mechanism. Neurochem Res 31: 509-514. Albert PR, Lembo P, Storring JM, Charest A, Saucier C. 1996. The 5‐HT1A receptor: Signaling, desensitization, and gene transcription. Neuropsychopharmacology 14: 19-25. Alex KD, Yavanian GJ, McFarlane HJ, Pluto CP, Pehek EA. 2005. Modulation of dopamine release by striatal 5‐HT2C receptors. Synapse 55: 242-251. Amlaiky N, Ramboz S, Boschert U, Plassat JL, Hen R. 1992. Isolation of a mouse ‘‘5HT1E‐like’’ serotonin receptor

expressed predominantly in hippocampus. J Biol Chem 267: 19761-19764. Anthony JP, Sexton TJ, Neumaier JF. 2000. Antidepressant‐ induced regulation of 5‐HT(1b) mRNA in rat dorsal raphe nucleus reverses rapidly after drug discontinuation. J Neurosci Res 61: 82-87. Arango V, Underwood MD, Boldrini M, Tamir H, Kassir SA, et al. 2001. Serotonin 1A receptors, serotonin transporter binding and serotonin transporter mRNA expression in the brainstem of depressed suicide victims. Neuropsychopharmacology 25: 892-903. Arango V, Underwood MD, Gubbi AV, Mann JJ. 1995. Localized alterations in pre‐ and postsynaptic serotonin binding sites in the ventrolateral prefrontal cortex of suicide victims. Brain Res 688: 121-133. Artigas F, Perez V, Alvarez E. 1994. Pindolol induces a rapid improvement of depressed patients with serotonin reuptake inhibitors. Arch Gen Psychiat 51: 248-251. Ase AR, Reader TA, Hen R, Riad M, Descarries L. 2000. Altered serotonin and dopamine metabolism in the CNS of serotonin 5‐HT(1A) or 5‐HT(1B) receptor knockout mice. J Neurochem 75: 2415-2426. Audenaert K, Laere Van K, Dumont F, Vervaet M, Goethals I, et al. 2003. Decreased 5‐HT2a receptor binding in patients with anorexia nervosa. J Nucl Med 44: 163-169. Aznar S, Qian Z, Shah R, Rahbek B, Knudsen GM. 2003. The 5‐HT1A serotonin receptor is located on calbindin‐ and parvalbumin‐containing neurons in the rat brain. Brain Res 959: 58-67. Bacon WL, Beck SG. 2000. 5‐Hydroxytryptamine7 receptor activation decreases slow afterhyperpolarization amplitude in CA3 hippocampal pyramidal cells. J Pharmacol Exp Ther 294: 672-679. Bai F, Yin T, Johnstone EM, Su C, Varga G, et al. 2004. Molecular cloning and pharmacological characterization of the guinea pig 5‐HT1E receptor. Eur J Pharmacol 484: 127-139. Baker LP, Nielson MD, Impey S, Hacker BM, Poser SW, et al. 1999. Regulation and immunohistochemical localization of

199

200

8

5‐Hydroxytryptamine in the central nervous system

betagamma‐stimulated adenylyl cyclases in mouse hippocampus. J Neurosci 19: 180-192. Bard JA, Zgombick J, Adham N, Vaysse P, Branchek TA, et al. 1993. Cloning of a novel human serotonin receptor (5‐HT7) positively linked to adenylate cyclase. J Biol Chem 268: 23422-23426. Barnes JM, Barnes NM, Costall B, Ironside JW, Naylor RJ. 1989a. Identification and characterisation of 5‐HT3 recognition sites in human brain tissue. J Neurochem 53: 1787-1793. Barnes JM, Barnes NM, Costall B, Naylor RJ, Tyers MB. 1989b. 5‐HT3 receptors mediate inhibition of acetylcholine release in cortical tissue. Nature 338: 762-763. Barnes JM, Costall B, Coughlan J, Domeney AM, Gerrard PA, et al. 1990. The effects of ondansetron, a 5‐HT3 receptor antagonist, on cognition in rodents and primates. Pharmacol Biochem Behav 35: 955-962. Barnes NM, Sharp T. 1999. A review of central 5‐HT receptors and their function. Neuropharmacology 38: 1083-1152. Beck SG, Pan, YZ, Akanwa AC, Kirby LG. 2004. Median and dorsal raphe neurons are not electrophysiologically identical. J Neurophysiol 91: 994-1005. Benloucif S, Keegan MJ, Galloway MP. 1993. Serotonin‐facilitated dopamine release in vivo‐pharmacological characterisation. J Pharmacol Exp Ther 265: 373-377. Bentley JC, Bourson A, Boess FG, Fone KCF, Marsden CA, et al. 1999. Investigation of stretching behaviour induced by the selective 5‐HT6 receptor antagonist, Ro 04‐6790, in rats. Brit J Pharmacol 126: 1537-1542. Berthouze M, Ayoub M, Russo O, Rivail L, Sicsic S, et al. 2005. Constitutive dimerization of human serotonin 5‐HT4 receptors in living cells. FEBS Lett 579: 2973-2980. Bianchi C, Siniscalchi A, Beani L. 1990. 5‐HT1A agonists increase and 5‐HT3 agonists decrease acetylcholine efflux from the cerebral cortex of freely‐moving guinea‐pigs. Brit J Pharmacol 101: 448-452. Blackburn TP, Minabe Y, Middlemiss DN, Shirayama Y, Hashimoto K, et al. 2002. Effect of acute and chronic administration of the selective 5‐HT2C receptor antagonist SB‐243213 on midbrain dopamine neurons in the rat: An in vivo extracellular single cell study. Synapse 46: 129-139. Blakely RD, Berson HE, Fremeau RTJ, Caron MG, Peek MM, et al. 1991. Cloning and expression of a functional serotonin transporter from rat brain. Nature 354: 66-70. Boess FG, Beroukhim R, Martin IL. 1995. Ultrastructure of the 5‐hydroxytryptamine(3) receptor. J Neurochem 64: 1401-1405. Bojarski AJ, Paluchowska MH, Duszynska B, Bugno R, Klodzinska A, et al. 2006. Structure–intrinsic activity relationship studies in the group of 1‐imido/amido substituted 4‐(4‐arylpiperazin‐1‐yl) cyclohexane derivatives; new,

potent 5‐HT1A receptor agents with anxiolytic‐like activity. BioorganMed Chem 14: 1391-1402. Bonaventure P, Schotte A, Cras P, Leysen JE. 1997. Autoradiographic mapping of 5‐HT1B‐ and 5‐HT1D receptors in human brain using [3H]alniditan, a new radioligand. Receptor Channel 5: 225-230. Boothman LJ, Allers KA, Rasmussen K, Sharp T. 2003. Evidence that central 5‐HT2A and 5‐HT2B/C receptors regulate 5‐HT cell firing in the dorsal raphe nucleus of the anaesthetised rat. Brit J Pharmacol 139: 998-1004. Boothman LJ, Mitchell SN, Sharp T. 2006. Investigation of the SSRI augmentation properties of 5‐HT2 receptor antagonists using in vivo microdialysis. Neuropharmacology 50: 726-732. Boschert U, Amara DA, Segu L, Hen R. 1994. The mouse 5‐hydroxytryptamine1B receptor is localized predominantly on axon terminals. Neuroscience 58: 167-182. Bosker FJ, Esseveldt van KE, Klompmakers AA, Westenberg HG. 1995. Chronic treatment with fluvoxamine by osmotic minipumps fails to induce persistent functional changes in central 5‐HT1A and 5‐HT1B receptors, as measured by in vivo microdialysis in dorsal hippocampus of conscious rats. Psychopharmacology 117: 358-363. Bouhelal R, Smounya L, Bockaert J. 1988. 5‐HT1B receptors are negatively coupled with adenylate cyclase in rat substantia nigra. Eur J Pharmacol 151: 189-196. Bourson A, Borroni E, Austin RH, Monsma FJ, Sleight AJ. 1995. Determination of the role of the 5‐ht6 receptor in the rat brain: A study using antisense oligonucleotides. J Pharmacol Exp Ther 274: 173-180. Boutrel B, Monaca C, Hen R, Hamon M, Adrien J. 2002. Involvement of 5‐HT1A receptors in homeostatic and stress‐ induced adaptive regulations of paradoxical sleep: Studies in 5‐HT1A knock‐out mice. J Neurosci 22: 4686-4692. Bramley JR, Sollars PJ, Pickard GE, Dudek FE. 2005. 5‐HT1B receptor‐mediated presynaptic inhibition of GABA release in the suprachiasmatic nucleus. J Neurophysiol 93: 3157-3164. Bruinvels AT, Landwehrmeyer B, Gustafson EL, Durkin MM, Mengod G, et al. 1994. Localization of 5‐HT1B, 5‐HT1Da, 5‐HT1E and 5‐HT1F, receptor messenger RNA in rodent and primate brain. Neuropharmacology 33: 367-386. Bruinvels AT, Palacios JM, Hoyer D. 1993. Autoradiographic characterisation and localisation of 5‐HT1D compared to 5‐HT1B binding sites in rat brain. N‐S Arch Pharmacol 347: 569-582. Bruning G, Liangos O. 1997. Transient expression of the serotonin transporter in the developing mouse thalamocortical system. Acta Histochem 99: 117-121. Brunner D, Buhot MC, Hen R, Hofer M. 1999. Anxiety, motor activation, and maternal–infant interactions in 5‐HT1B knockout mice. Behav Neurosci 113: 587-601.

5‐Hydroxytryptamine in the central nervous system Burnet PWJ, Eastwood SL, Harrison PJ. 1996. 5‐HT1A and 5‐HT2A receptor mRNAs and binding site densities are differentially altered in schizophrenia. Neuropsychopharmacology 15: 442-455. Burnet PWJ, Eastwood SL, Lacey K, Harrison PJ. 1995. The distribution of 5HT1A and 5‐HT2A receptor mRNA in human brain. Brain Res 676: 157-168. Burns CM, Chu H, Rueter SM, Hutchinson LK, Canton H, et al. 1997. Regulation of serotonin‐2C receptor G‐protein coupling by RNA editing. Nature 387: 303-308. Cai X, Gu Z, Zhong P, Ren Y, Yan Z. 2002. Serotonin 5‐HT1A receptors regulate AMPA receptor channels through inhibiting Ca2þ/calmodulin‐dependent kinase II in prefrontal cortical pyramidal neurones. J Biol Chem 277: 3655336562. Carter OL, Burr DC, Pettigrew JD, Wallis GM, Hasler F, et al. 2005. Using psilocybin to investigate the relationship between attention, working memory, and the serotonin 1A and 2A receptors. J Cognitive Neurosci 17: 1497-1508. Caspi A, Sugden K, Moffitt TE, Taylor A, Craig IW, et al. 2003. Influence of life stress on depression: Moderation by a polymorphism in the 5‐HTT gene. Science 301: 386-389. Cassel JC, Jeltsch H, Neufang B, Lauth D, Szabo B, et al. 1995. Downregulation of muscarinic‐ and 5‐HT1B‐mediated modulation of [3H]acetylcholine release in hippocampal slices of rats with fimbria–fornix lesions and intrahippocampal grafts of septal origin. Brain Res 704: 153-166. Castro ME, Pascual J, Romon T, del Arco C, del Olmo E, et al. 1997. Differential distribution of [3H]sumatriptan binding sites (5‐HT1B, 5‐HT1D and 5‐HT1F receptors) in human brain: Focus on brainstem and spinal cord. Neuropharmacology 36: 535-542. Chapin EM, Andrade R. 2001. A 5‐HT7 receptor‐mediated depolarization in the anterodorsal thalamus. ii. involvement of the hyperpolarization‐activated current Ih. J Pharmacol Exp Ther 297: 403-409. Chapin EM, Haj‐Dahmane S, Torres G, Andrade R. 2002. The 5‐HT(4) receptor‐induced depolarization in rat hippocampal neurons is mediated by cAMP but is independent of I(h). Neurosci Lett 324: 1-4. Cheetham SC, Crompton MR, Katona CLE, Horton RW. 1990. Brain 5‐HT1 binding sites in depressed suicides. Psychopharmacology 102: 544-548. Chen NH, Reith ME. 1995. Monoamine interactions measured by microdialysis in the ventral tegmental area of rats treated systemically with (9)‐8‐hydroxy‐2‐(di‐n‐propylamino)tetralin. J Neurochem 64: 1585-1597. Clark MS, Sexton TJ, McClain M, Root D, Kohen R, et al. 2002. Overexpression of 5‐HT1B receptor in dorsal raphe nucleus using herpes simplex virus gene transfer increases anxiety behavior after inescapable stress. J Neurosci 22: 4550-4562.

8

Clarke WP, Yocca FD, Maayani S. 1996. Lack of 5‐hydroxytryptamine1A‐mediated inhibition of adenylyl cyclase in dorsal raphe of male and female rats. J Pharmacol Exp Ther 277: 1259-1266. Cohen H. 2005. Anxiolytic effect and memory improvement in rats by antisense oligodeoxynucleotide to 5‐hydroxytryptamine‐2A precursor protein. Depress Anxiety 22: 84-93. Compan V, Zhou M, Grailhe R, Gazzara RA, Martin R, et al. 2004. Attenuated response to stress and novelty and hypersensitivity to seizures in 5‐HT4 receptor knock‐out mice. J Neurosci 24: 412-419. Conn PJ, Sanders‐Bush E. 1984. Selective 5‐HT2 antagonists inhibit serotonin stimulated phosphatidyl inositol metabolism in cerebral cortex. Neuropharmacology 23: 993-996. Consolo S, Arnaboldi S, Giorgi S, Russi G, Ladinsky H. 1994. 5‐HT4 receptor stimulation facilitates acetylcholine release in rat frontal cortex. NeuroReport 5: 1230-1232. Corbett DF, Heightman TD, Moss SF, Bromidge SM, Coggon SA, et al. 2005. Discovery of a potent and selective 5‐ht5A receptor antagonist by high‐throughput chemistry. Bioorgan Med Chem Lett 15: 4014-4018. Cornea‐Hebert V, Watkins KC, Roth BL, Kroeze WK, Gaudreau P, et al. 2002. Similar Ultrastructural distribution of the 5‐HT2A serotonin receptor and microtubule‐associated protein MAP1A in cortical dendrites of adult rat. Neuroscience 113: 23-35. Cortes R, Soriano E, Pazos A, Probst A, Palacios JM. 1988. Autoradiography of antidepressant binding sites in the human brain: Localization using [3H]‐imipramine and [3H]‐paroxetine. Neuroscience 27: 473-496. Costall B, Kelly ME, Naylor RJ, Onaivi ES, Tyers MB. 1989. Neuroanatomical sites of action of 5‐HT3 receptor agonists and antagonists for alteration of aversive behaviour in the mouse. Brit J Pharmacol 96: 325-332. Crabbe JC, Phillips TJ, Feller DJ, Hen R, Wenger CD, et al. 1996. Elevated alcohol consumption in null mutant mice lacking 5‐HT1B serotonin receptors. Nat Genet 14: 98-101. D’Amato RJ, Blue ME, Largent BL, Lynch DR, Ledbetter DJ, et al. 1987. Ontogeny of the serotonergic projection to rat neocortex: Transient expression of a dense innervation to primary sensory areas. PNAS 84: 4322-4326. Dahlstro¨m A, Fuxe K. 1964. Evidence for the existence of monoamine containing neurons in the central nervous system. I. Demonstration of monoamines in cell bodies of brain neurons. Acta physiol Scand 62 (Suppl 232): 1-55. Darmon M, Langlois X, Suffisseau L, C‐M, Fattaccini Hamon M. 1998. Differential membrane targeting and pharmacological characterisation of chimeras of rat serotonin 5‐HT1A and 5‐HT1B receptors expressed in epithelial LLC‐PK1 cells. J Neurochem 71: 2294-2303.

201

202

8

5‐Hydroxytryptamine in the central nervous system

Davies PA, Pistis M, Hanna MC, Peters JA, Lambert JL, et al. 1999. The 5‐HT3B subunit is a major determinant of serotonin‐receptor function. Nature 397: 359-363. Dawes MA, Johnson BA, Ait‐Daoud N, Ma JZ, Cornelius JR. 2005. A prospective, open‐label trial of ondansetron in adolescents with alcohol dependence. Addict Behav 30: 1077-1085. Dawson LA, Nguyen HQ, Li P. 2001. The 5‐HT6 receptor antagonist SB‐271046 selectively enhances excitatory neurotransmission in the rat frontal cortex and hippocampus. Neuropsychopharmacology 25: 662-668. de Boer SF, Koolhass JM. 2005. 5‐HT1A and 5‐HT1B receptor agonists and aggression: A pharmacological challenge of the serotonin deficiency hypothesis. Eur J Pharmacol 526: 125-139. De Deurwaerdere P, Spampinato U. 1999. Role of serotonin2A and serotonin2B/2C receptor subtypes in the control of accumbal and striatal dopamine release elicited in vivo by dorsal raphe nucleus electrical stimulation. J Neurochem 73: 1033-1042. De Deurwaerdere P, Stinus L, Spampinato U. 1998. Opposite change of in vivo dopamine release in the rat nucleus accumbens and striatum that follows electrical stimulation of dorsal raphe nucleus: Role of 5‐HT3 receptors. J Neurosci 18: 6528-6538. Den Boer JA, Bosker FJ, Meesters Y. 2006. Clinical efficacy of agomelatine in depression: The evidence. Int Clin Psychopharmacol 21: S21-S24. Di Giovanni G, De Deurwaerdere P, Di Mascio M, Di Matteo V, Esposito E, et al. 1999. Selective blockade of serotonin‐ 2C/2B receptors enhances mesolimbic and mesostriatal dopaminergic function: A combined in vivo electrophysiological and microdialysis study. Neuroscience 91: 587-597. Di Matteo V, Di Giovanni G, Di Mascio M, Esposito E. 1999. SB 242 084, a selective serotonin2C receptor antagonist, increases dopaminergic transmission in the mesolimbic system. Neuropharmacology 38: 1195-1205. Dı´az‐Mataix L, Scorza MC, Bortolozzi A, Toth M, Celada P, et al. 2005. Involvement of 5‐HT1A receptors in prefrontal cortex in the modulation of dopaminergic activity: Role in atypical antipsychotic action. J Neurosci 25: 1083110843. Diez‐Ariza M, Garcia‐Alloza M, Lasheras B, Del Rio J, Ramirez MJ. 2002. GABA(A) receptor antagonists enhance cortical acetylcholine release induced by 5‐HT(3) receptor blockade in freely moving rats. Brain Res 956: 81-85. Done CJ, Sharp T. 1994. Biochemical evidence for the regulation of central noradrenergic activity by 5‐HT1A and 5‐HT2 receptors: Microdialysis studies in the awake and anaesthetized rat. Neuropharmacology 33: 411-421. Du Pen S, Scuderi P, Wetchler B, Sung YF, Mingus M, et al. 1992. Ondansetron in the treatment of postoperative

nausea and vomiting in ambulatory outpatients: A dose‐comparative, stratified, multicentre study. Eur J Anaesthesiol 9: 55-62. Dubin AE, Huvar R, D’Andrea MR, Pyati J, Zhu JY, et al. 1999. The pharmacological and functional characteristics of the serotonin 5‐HT3A receptor are specifically modified by a 5‐HT3B receptor subunit. J Biol Chem 274: 3079930810. Dukat M, Smith C, Herrick‐Davis K, Teitler M, Glennon RA. 2004. Binding of tryptamine analogs at h5‐HT1E receptors: A structure–affinity investigation. Bioorgan Med Chem 12: 2545-2552. Duncan MJ, Grear KE, Hoskins MA. 2004. Aging and SB‐ 269970‐A, a selective 5‐HT7 receptor antagonist, attenuate circadian phase advances induced by microinjections of serotonergic drugs in the hamster dorsal raphe nucleus. Brain Res 1008: 40-48. Duncan MJ, Short J, Wheeler DL. 1999. Comparison of the effects of aging on 5‐HT7 and 5‐HT1A receptors in discrete regions of the circadian timing system in hamsters. Brain Res 829: 39-45. Duxon MS, Flanigan TP, Reavley AC, Baxter GS, Blackburn TP, et al. 1997a. Evidence for the expression of the 5‐hydroxytryptamine2B receptor protein in the rat central nervous system. Neuroscience 76: 323-329. Duxon MS, Kennett GA, Lightowler S, Blackburn TP, Fone KC. 1997b. Activation of 5‐HT2B receptors in the medial amygdala causes anxiolysis in the social interaction test in the rat. Neuropharmacology 36: 601-608. East SZ, Burnet PW, Kerwin RW, Harrison PJ. 2002b. An RT‐ PCR study of 5‐HT(6) and 5‐HT(7) receptor mRNAs in the hippocampal formation and prefrontal cortex in schizophrenia. Schizoph Res 57: 15-26. East SZ, Burnet PWJ, Harrison PJ. 1999. Expression of the 5‐HT7 receptor in schizophrenia. Schizophr Res 36: 71. East SZ, Burnet PWJ, Leslie RA, Roberts JC, Harrison PJ. 2002a. 5‐HT6 receptor binding sites in schizophrenia and following antipsychotic drug administration: Autoradiographic studies with [125I]SB‐258585. Synapse 45: 191-199. Egashira N, Yano A, Ishigami N, Mishima K, Iwasaki K, et al. 2006. Investigation of mechanisms mediating 8‐OH‐DPAT‐ induced impairment of spatial memory: Involvement of 5‐ HT1A receptors in the dorsal hippocampus in rats. Brain Res 1069: 54-62. Fa¨rber L, Stratz T, Bruckle W, Spatz M, Pongratz D, et al. 2000. Efficacy and tolerability of tropisetron in primary fibromyalgia: A highly selective and competitive 5‐HT3 receptor antagonist. Scand J Rheumatol 113: 49-54. Federman AD, Conklin BR, Schrader KA, Reed RR, Bourne HR. 1992. Hormonal stimulation of adenylyl cyclase through Gi‐protein beta gamma subunits. Nature 356: 159-161.

5‐Hydroxytryptamine in the central nervous system Ferris CF, Melloni RH, Koppel G, Perry KW, Fuller RW, et al. 1997. Vasopressin/serotonin interaction in the anterior hypothalamus control aggressive behavior in golden hamsters. J Neurosci 17: 4331-4340. Fletcher A, Forster EA, Bill DJ, Brown G, Cliffe IA, et al. 1996. Electrophysiological, biochemical, neurohormonal and behavioural studies with WAY 100635, a potent, selective and silent 5‐HT1A receptor antagonist. Behav Brain Res 73: 337-353. Foguet M, Hoyer D, Pardo LA, Parekh A, Kluxen FW, et al. 1992. Cloning and functional characterization of the rat stomach fundus serotonin receptor. EMBO J 11: 34813487. Francken BJB, Jurzak M, Vanhauwe JFM, Luyten WHML, Leysen JE. 1998. The human 5‐ht5A receptor couples to Gi/Go proteins and inhibits adenylate cyclase in HEK 293 cells. Eur J Pharmacol 361: 299-309. Frankle WG, Narendran R, Huang Y, Hwang DR, Lombardo I, et al. 2005. Serotonin Transporter availability in patients with schizophrenia: A positron emission tomography imaging study with [11C]DASB. Biol Psychiat 57: 1510-1516. Frederick JA, Meador‐Woodruff JH. 1999. Effects of clozapine and haloperidol on 5‐HT6 receptor mRNA levels in rat brain. Schizophr Res 38: 7-12. Fujita M, Shimada S, Maeno H, Nishimura T, Tohyama M. 1993. Cellular localization of serotonin transporter mRNA in the rat brain. Neurosci Lett 162: 59-62. Gannon RL, Millan MJ. 2006. Serotonin1A autoreceptor activation by S 15535 enhances circadian activity rhythms in hamsters: Evaluation of potential interactions with serotonin2A and serotonin2C receptors. Neuroscience 137: 287-299. Garabette ML, Martin KF, Redfern PH. 2000. Circadian variation in the activity of the 5‐HT1B autoreceptor in the region of the suprachiasmatic nucleus, measured by microdialysis in the conscious freely‐moving rat. Brit J Pharmacol 131: 1569-1576. Ge J, Barnes NM. 1996. 5‐HT4 receptor mediated modulation of 5‐HT release in the rat hippocampus in vivo. Brit J Pharmacol 17: 1474-1480. Gerald C, Adham A, Kao HT, Olsen MA, Laz TM, et al. 1995. The 5‐HT4 receptor: Molecular cloning and pharmacological characterisation of two splice variants. EMBO J 14: 2806-2815. Gerard C, Martres MP, Lefevre K, Miquel MC, Verge D, et al. 1997. Immuno‐localization of serotonin 5‐HT6 receptor‐ like material in the rat central nervous system. Brain Res 746: 207-219. Goa KL, Ward A. 1986. Buspirone. A preliminary review of its pharmacological properties and therapeutic efficacy as an anxiolytic. Drugs 32: 114-129.

8

Goaillard JM, Vincent P. 2002. Serotonin suppresses the slow afterhyperpolarization in rat intralaminar and midline thalamic neurones by activating 5‐HT7 receptors. J Physiol 541: 453-465. Gobert A, Rivet JM, Cistarelli L, Millan MJ. 1997. Potentiation of the fluoxetine‐induced increase in dialysate levels of serotonin (5‐HT) in the frontal cortex of freely moving rats by combined blockade of 5‐HT1A and 5‐HT1B receptors with WAY 100,635 and GR 127,935. J Neurochem 68: 1159-1163. Graf M, Jakus R, Kantor S, Levay G, Bagdy G. 2004. Selective 5‐HT1A and 5‐HT7 antagonists decrease epileptic activity in the WAG/Rij rat model of absence epilepsy. Neurosci Lett 359: 45-48. Grailhe R, Grabtree GW, Hen R. 2001. Human 5‐HT5 receptors: The 5‐HT5A receptor is functional but the 5‐HT5B receptor was lost during mammalian evolution. Eur J Pharmacol 418: 157-167. Grailhe R, Waeber C, Dulawa SC, Hornung JP, Zhuang X, et al. 1999. Increased exploratory activity and altered response to LSD in mice lacking the 5‐HT5A receptor. Neuron 22: 581-591. Green GM, Scarth J, Dickenson A. 2000. An excitatory role for 5‐HT in spinal inflammatory nociceptive transmission; state‐dependent actions via dorsal horn 5‐HT3 receptors in the anaesthetized rat. Pain 89: 81-88. Greenberg BD, Tolliver TJ, Huang SJ, Li Q, Bengel D, et al. 1999. Genetic variation in the serotonin transporter promoter region affects serotonin uptake in human blood platelets. Am J Med Genet 88: 83-87. Grimaldi B, Bonnin A, Fillion MP, Prudhomme N, Fillion G. 1999. 5‐HT moduline: A novel endogenous peptide involved in the control of anxiety. Neuroscience 93: 1223-1225. Groenink L, Mos J, der Gugten Van J, Olivier B. 1996. The 5‐HT1A receptor is not involved in emotional stress‐induced rises in stress hormones. Pharmacol Biochem Behav 55: 303-308. Gurevich EV, Joyce JN. 1997. Alterations in the cortical serotonergic system in schizophrenia: A postmortem study. Biol Psychiat 42: 529-545. Gurevich I, Englander MT, Adlersberg M, Siegal NB, Schmauss C. 2002. Modulation of serotonin 2C receptor editing by sustained changes in serotonergic neurotransmission. J Neurosci 22: 10529-10532. Guscott M, Bristow LJ, Hadingham K, Rosahl TW, Beera MS, et al. 2005. Genetic knockout and pharmacological blockade studies of the 5‐HT7 receptor suggest therapeutic potential in depression. Neuropharmacology 48: 492-502. Hagan JJ, Price GW, Jeffrey P, Deeks NJ, Stean T, et al. 2000. Characterization of SB‐269970‐A, a selective 5‐HT(7) receptor antagonist. Brit J Pharmacol 130: 539-548.

203

204

8

5‐Hydroxytryptamine in the central nervous system

Hamblin MW, Metcalf MA. 1991. Primary structure and functional characterization of a human 5‐HT1D‐type serotonin receptor. Mol Pharmacol 40: 143-148. Hamon M, Doucet E, Lefe`vre K, Miquel MC, Lanfumey L, et al. 1999. Antibodies antisense oligonucleotide for probing the distribution and putative functions of central 5‐HT6 receptors. Neuropsychopharmacology 21: 68S-76S. Hariri AR, Mattay VS, Tessitore A, Kolachana B, Fera F, et al. 2002. Serotonin transporter genetic variation and the response of the human amygdala. Science 297: 400-403. Harsing LG, Prauda I, Barkoczy J, Matyus P, Juranyi Z. 2004. A 5‐HT7 heteroreceptor‐mediated inhibition of [3H]serotonin release in raphe nuclei slices of the rat: Evidence for a serotonergic–glutamatergic interaction. Neurochem Res 29: 1487-1497. Hartig PR, Hoyer D, Humphrey PPA, Martin GR. 1996. Alignment of receptor nomenclature with the human genome: Classification of 5‐HT1B and 5‐HT1D receptor subtypes. Trends Pharmacol Sci 17: 103-105. Harvey JA. 2003. Role of the serotonin 5‐HT2A receptor in learning. Learn Memory 10: 355-362. Hedlund PB, Huitron‐Resendiz S, Henriksen SJ, Sutcliffe JG. 2005. 5‐HT7 receptor inhibition and inactivation induce antidepressant‐like behavior and sleep pattern. Biol Psychiat 58: 831-837. Hedlund PB, Kelly L, Mazur C, Lovenberg T, Sutcliffe JG, et al. 2004. 8‐OH‐DPAT acts on both 5‐HT1A and 5‐HT7 receptors to induce hypothermia in rodents. Eur J Pharmacol 487: 125-132. Heidmann DEA, Metcalf MA, Kohen R, Hamblin MW. 1997. Four 5 hydroxytryptamine7 (5‐HT7) receptor isoforms in human and rat produced by alternative splicing: Species differences due to altered intron–exon organization. J Neurochem 68: 1372-1381. Heidmann DEA, Szot P, Kohen R, Hamblin MW. 1998. Function and distribution of three rat 5‐hydroxytryptamine7 (5‐HT7) receptor isoforms produced by alternative splicing. Neuropharmacology 37: 1621-1632. Helsley S, Siegel TL, Fiorella D, Rabin RA, Winter JC. 1998. WAY‐100635 reverses 8‐OH‐DPAT‐induced performance impairment in the radial maze. Prog Neuropsychopharmacol Biol Psychiat 22: 1179-1184. Hensler JG. 2006. Serotonergic modulation of the limbic system. Neurosci Biobehav Rev 30: 203-214. Hirst WD, Abrahamsen B, Blaney FE, Calver AR, Aloj L, et al. 2003. Differences in the central nervous system distribution and pharmacology of the mouse 5‐hydroxytryptamine‐6 receptor compared with rat and human receptors investigated by radioligand binding, site‐directed mutagenesis, and molecular modeling. Mol Pharmacol 64: 1295-1308.

Hjorth S, Sharp T. 1991. Effect of the 5‐HT1A receptor agonist 8‐OH‐DPAT on the release of 5‐HT in dorsal and median raphe innervated rat brain regions as measured by in vivo microdialysis. Life Sci 48: 1779-1786. Hornung JP. 2003. The human raphe nuclei and the serotonergic system. J Chem Neuroanatomy 26: 331-343. Hou M, Kanje M, Longmore J, Tajti J, Uddman R, et al. 2001. 5‐HT1B and 5‐HT1D receptors in the human trigeminal ganglion: Co‐localization with calcitonin gene‐related peptide, substance P and nitric oxide synthase. Brain Res 909: 112-120. Hoyer D, Clarke DE, Fozard JR, Hartig PR, Martin GR, et al. 1994. International Union of Pharmacology classification of receptors for 5‐hydroxytryptamine (serotonin). Pharmacol Rev 46: 157-204. Hu XZ, Lipsky RH, Zhu G, Akhtar LA, Taubman J, et al. 2006. Serotonin transporter promoter gain‐of‐function genotypes are linked to obsessive–compulsive disorder. Am J Human Genet 78: 815-826. Huang GJ, Herbert J. 2005. The role of 5‐HT1A receptors in the proliferation and survival of progenitor cells in the dentate gyrus of the adult hippocampus and their regulation by corticoids. Neuroscience 135: 803-813. Hurley PT, McMahon RA, Fanning P, O’Boyle KM, Rogers M, et al. 1998. Functional coupling of a recombinant human 5‐HT5A receptor to G‐proteins in HEK‐293 cells. Brit J Pharmacol 124: 1238-1244. Ichise M, Vines DC, Gura T, Anderson GM, Suomi SJ, et al. 2006. Effects of early life stress on [11C]DASB positron emission tomography imaging of serotonin transporters in adolescent peer‐ and mother‐reared rhesus monkeys. J Neurosci 26: 4638-4643. Idzikowski C, Mills FJ, James RJ. 1991. A dose–response study examining the effects of ritanserin on human slow wave sleep. Brit J Clin Pharmacol 31: 193-196. Ikeda M, Yasui M, Fukunaga H, Seshimo I, Takayama O, et al. 2005. Clinical usefulness of oral granisetron hydrochloride for alleviation of delayed nausea and vomiting induced by CPT‐11. Eur J Cancer Care 14: 435-439. Ikemoto K, Nishimura A, Okado N, Mikuni M, Nishi K, et al. 2000. Human midbrain dopamine neurons express serotonin 2A receptor: An immunohistochemical demonstration. Brain Res 853: 377-380. Inui K, Egashira N, Mishima K, Yano A, Matsumoto Y, et al. 2004. The serotonin1A receptor agonist 8‐OH‐DPAT reverses D9‐tetrahydrocannabinol‐induced impairment of spatial memory and reduction of acetylcholine release in the dorsal hippocampus in rats. Neurotox Res 6: 153-158. Jakab R, Goldman‐Rakic PS. 1998. 5‐Hydroxytryptamine2A serotonin receptors in the primate cerebral cortex: Possible

5‐Hydroxytryptamine in the central nervous system site of action of hallucinogenic and antipsychotic drugs in pyramidal cell apical dendrites. PNAS 95: 735-740. Johnson KW, Nelson DL, Dieckman DK, Wainscott DB, Lucaites VL, et al. 2003. Neurogenic dural protein extravasation induced by meta‐chlorophenylpiperazine (mCPP) involves nitric oxide and 5‐HT2B receptor activation. Cephalalgia 23: 117-123. Jorgensen H, Riis M, Knigge U, Kjaer A, Warberg J. 2003. Serotonin receptors involved in vasopressin and oxytocin secretion. J Neuroendocrinol 15: 242-249. Julius D, Huang KN, Livelli TJ, Axel R, Jessell TM. 1990. The 5HT2 receptor defines a family of structurally distinct but functionally conserved serotonin receptors. PNAS 87: 928-932. Julius D, Mac Dermott AB, Axel R, Jessell TM. 1988. Molecular characterisation of a functional cDNA encoding the serotonin1C receptor. Science 244: 558-564. Kantor S, Jakus R, Balogh B, Benko A, Bagdy G. 2004. Increased wakefulness, motor activity and decreased theta activity after blockade of the 5‐HT2B receptor by the subtype‐selective antagonist SB‐215505. Brit J Pharmacol 142: 1332-1342. Kantor S, Jakus R, Molnar E, Gyongyosi N, Toth A, et al. 2005. Despite similar anxiolytic potential, the 5‐hydroxytryptamine 2C receptor antagonist SB‐242084 [6‐chloro‐5‐ methyl‐1‐[2‐(2‐methylpyrid‐3‐yloxy)‐pyrid‐5‐yl carbamoyl] indoline] and chlordiazepoxide produced differential effects on electroencephalogram power spectra. J Pharmacol Exp Ther 315: 921-930. Kapczinski F, Lima MS, Souza JS, Cunha A, Schmitt R. 2003. Antidepressants for generalized anxiety disorder. The Cochrane Database of Systematic Reviews Issue 2: Article No.: CD003592. Kelley SP, Bratt AM, Hodge CW. 2003. Targeted gene deletion of the 5‐HT3A receptor subunit produces an anxiolytic phenotype in mice. Eur J Pharmacol 461: 19-25. Kennett GA, Ainsworth K, Trail B, Blackburn TP. 1997a. BW 723C86, a 5‐HT2B receptor agonist, causes hyperphagia and reduced grooming in rats. Neuropharmacology 36: 233-239. Kennett GA, Bright F, Trail B, Blackburn TP, Sanger GJ. 1997c. Anxiolytic‐like actions of the selective 5‐HT4 receptor antagonists, SB204070A and SB207266A in rats. Neuropharmacology 36: 707-712. Kennett GA, Trail B, Bright F. 1998. Anxiolytic‐like actions of BW 723C86 in the rat Vogel conflict test are 5‐HT2B receptor mediated. Neuropharmacology 37: 1603-1610. Kennett GA, Wood MD, Bright F, Cilia J, Piper DC, et al. 1996. In vitro and in vivo profile of SB 206553, a potent

8

5‐HT2C:5‐HT2B receptor antagonist with anxiolytic‐like properties. Brit J Pharmacol 117: 427-434. Kennett GA, Wood MD, Bright F, Trail B, Riley G, et al. 1997b. SB 242084, a selective and brain penetrant 5‐HT2C receptor antagonist. Neuropharmacology 36: 609-620. Kepe V, Barrio JR, Huang SC, Ercoli L, Siddarth P, et al. 2006. Serotonin 1A receptors in the living brain of Alzheimer’s disease patients. PNAS 103: 702-707. Kia HK, Brisorgueil MJ, Daval G, Langlois X, Hamon M, et al. 1996a. Serotonin 5‐HT1A receptors expressed by a subpopulation of cholinergic neurons in the rat medial septum and diagonal band of Broca – a double immunocytochemical study. Neuroscience 74: 143-154. Kia HK, Brisorgueil MJ, Hamon M, Calas A, Verge D. 1996b. Ultrastructural localization of 5‐hydroxytryptamine1A receptors in rat brain. J Neurosci Res 46: 697-708. Kilic F, Rudnick G. 2000. Oligomerization of serotonin transporter and its functional consequences. PNAS 97: 31063111. King MV, Sleight AJ, Woolley ML, Topham IA, Marsden CA, et al. 2004. 5‐HT6 receptor antagonists reverse delay‐ dependent deficits in novel object discrimination by enhancing consolidation – an effect sensitive to NMDA receptor antagonism. Neuropharmacology 47: 195-204. Kobilka BK, Frielle T, Collins S, Yang‐Feng T, Kobilka TS, et al. 1987. An intronless gene encoding a potential member of the family of receptors coupled to guanine nucleotide regulatory proteins. Nature 329: 75-77. Koyama S, Matsumoto N, Kubo C, Akaike N. 2000. Presynaptic 5‐HT3 receptor‐mediated modulation of synaptic GABA release in the mechanically dissociated rat amygdala neurons. J Physiol 529: 373-383. Krobert KA, Bach T, Syversveen T, Kvingedal AM, Levy FO. 2001. The cloned human 5‐HT7 receptor splice variants: A comparative characterization of their pharmacology, function and distribution. N‐S Arch Pharmacol 363: 620-632. Lacroix LP, Dawson LA, Hagan JJ, Heidbreder CA. 2004. 5‐HT6 receptor antagonist SB‐271046 enhances extracellular levels of monoamines in the rat medial prefrontal cortex. Synapse 51: 158-164. Laruelle M, Vanisberg MA, Maloteaux JM. 1988. Regional and subcellular localization in human brain of [3H]paroxetine binding, a marker of serotonin uptake sites. Biol Psychiat 24: 299-309. Lawrence JA, Charters AR, Butcher SP, Kelly JS, Olverman HJ. 1995a. Recognition of 5‐HT transporter by antipeptide antibodies. Biochem Soc Trans 23: 473. Lawrence JA, Charters AR, Butcher SP, Kelly JS, Olverman HJ. 1995b. 5‐HT transporter antibodies as a tool in serotonergic synaptosomal isolation. Biochem Soc Trans 23: 1155.

205

206

8

5‐Hydroxytryptamine in the central nervous system

Lebrand C, Cases O, Aldebrecht C, Doye A, Alvarez C, et al. 1996. Transient uptake and storage of serotonin in developing thalamic neurons. Neuron 17: 823-835. Lelong V, Dauphin F, Boulouard M. 2001. RS 67333 and D‐cycloserine accelerate learning acquisition in the rat. Neuropharmacology 41: 517-522. Leng A, Ouagazzal A, Feldon J, Higgins GA. 2003. Effect of the 5‐HT6 receptor antagonists Ro 04‐6790 and Ro 65‐7199 on latent inhibition and prepulse inhibition in the rat: Comparison to clozapine. Pharmacol Biochem Behav 75: 281-288. Lesch KP, Bengel D, Heils A, Sabol SZ, Greenberg BD, et al. 1996. Association of anxiety‐related traits with a polymorphism in the serotonin transporter gene regulatory region. Science 274: 1527-1531. Levy FO, Gudermann T, Birnbaumer M, Kaumann AJ, Birnbaumer L. 1992. Molecular cloning of a human gene (S31) encoding a novel serotonin receptor mediating inhibition of adenylyl cyclase. FEBS Lett 296: 201-206. Lezoualc’h F, Robert SJ. 2003. The serotonin 5‐HT4 receptor and the amyloid precursor protein processing. Exp Gerontol 38: 159-166. Li Q, Holmes A, Ma L, de Kar Van LD, Garcia F, et al. 2004. Medial hypothalamic 5‐hydroxytryptamine (5‐HT)1A receptors regulate neuroendocrine responses to stress and exploratory locomotor activity: Application of recombinant adenovirus containing 5‐HT1A sequences. J Neurosci 24: 10868-10877. Lin D, Parsons LH. 2002. Anxiogenic‐like effect of serotonin (1B) receptor stimulation in the rat elevated plus‐maze. Pharmacol Biochem Behav 71: 581-587. Lindner MD, Hodges DB, Hogan JB, Orie AF, Corsa JA, et al. 2003. An assessment of the effects of serotonin 6 (5‐HT6) receptor antagonists in rodent models of learning. J Pharmacol Exp Ther 307: 682-691. Lira A, Zhou M, Castanon N, Ansorge MS, Gordon JA, et al. 2003. Altered depression‐related behaviors and functional changes in the dorsal raphe nucleus of serotonin transporter‐deficient mice. Biol Psychiat 54: 960-971. Little KY, Mclaughlin DP, Zhang L, Livermore CS, Dalack GW, et al. 1998. Cocaine, ethanol, and genotype effects on human midbrain serotonin transporter binding sites and mRNA levels. Am J Psychiat 155: 207-213. Lopez‐Gimenez JF, Mengod G, Palacios JM, Vilaro MT. 1997. Selective visualisation of rat brain 5‐HT2A receptors by autoradiography with [3H]MDL 100 907. N‐S Arch Pharmacol 356: 446-454. Lowther S, De Paermentier F, Cheetham SC, Crompton MR, Katona CL, et al. 1997. 5‐HT1A receptor binding sites in post‐mortem brain samples from depressed suicides and controls. J Affect Disorders 42: 199-207.

Lucaites VL, Krushinski JH, Schaus JM, Audia JE, Nelson DL. 2005. [3H]LY334370, a novel radioligand for the 5‐HT1F receptor. II. Autoradiographic localization in rat, guinea‐ pig, monkey and human brain. N‐S Arch Pharmacol 371: 178-184. Lucas G, Compan V, Charnay Y, Neve RL, Nestler EJ, et al. 2005. Frontocortical 5‐HT4 receptors exert positive feedback on serotonergic activity: Viral transfections, subacute and chronic treatments with 5‐HT4 agonists. Biol Psychiat 57: 918-925. Lucas G, Spampinato U. 2000. Role of striatal serotonin2A and serotonin2C receptor subtypes in the control of in vivo dopamine outflow in the rat striatum. J Neurochem 74: 693-701. Lucki I, Singh A, Kreiss DS. 1994. Antidepressant‐like behavioral effects of serotonin receptor agonists. Neurosci Biobehav Rev 18: 85-95. Ma QP. 2001. Co‐localization of 5‐HT1B/1D/1F receptors and glutamate in trigeminal ganglia in rats. NeuroReport 12: 1589-1591. Machu TK, Harris RA. 1994. Alcohols and anesthetics enhance the function of 5‐hydroxytryptamine3 receptors expressed in Xenopus laevis oocytes. J Pharmacol Exp Ther 271: 898-905. Malberg JE, Eisch AJ, Nestler EJ, Duman RS. 2000. Chronic antidepressant treatment increases neurogenesis in adult rat hippocampus. J Neurosci 20: 9104-9110. Malison RT, Price LH, Berman R, Dyck van CH, Pelton GH, et al. 1998. Reduced brain serotonin transporter availability in major depression as measured by [123I]‐2 beta‐carbomethoxy‐3 beta‐(4‐iodophenyl)tropane and single photon emission computed tomography. Biol Psychiat 44: 10901098. Martin KF, Hannon S, Phillips I, Heal DJ. 1992. Opposing roles for 5‐HT1B and 5‐HT3 receptors in the control of 5‐HT release in rat hippocampus in vivo. Brit J Pharmacol 106: 139-142. Martin‐Ruiz R, Puig MV, Celada P, Shapiro DA, Roth BL, et al. 2001. Control of serotonergic function in medial prefrontal cortex by serotonin‐2A receptors through a glutamate dependent mechanism. J Neurosci 21: 9856-9866. Masson J, Sagne C, Hamon M, El Mestikawy S. 1999. Neurotransmitter transporters in the central nervous system. Pharmacol Rev 51: 439-464. Massot O, Rousselle JC, Fillion MP, Grimaldi B, Cloez‐ Tayarani I, et al. 1996. 5‐hydroxytryptamine‐moduline, a new endogenous cerebral tetrapeptide, controls the serotonergic activity via its specific interaction with 5‐ hydroxytryptamine1B/1D receptors. Mol Pharmacol 50: 752-762.

5‐Hydroxytryptamine in the central nervous system Matthes H, Boschert U, Amlaiky N, Grailhe R, Plassat JL, et al. 1993. Mouse 5‐hydroxytryptamine 5A and 5‐hydroxytryptamine 5B receptors define a new family of serotonin receptors: Cloning, functional expression, and chromosomal localization. Mol Pharmacol 43: 313-319. Maura G, Marcoli M, Tortarolo M, Andrioli GC, Raiteri M. 1998. Glutamate release in human cerebral cortex and its modulation by 5‐hydroxytryptamine acting at h5‐HT1D receptors. Brit J Pharmacol 123: 45-50. McAllister G, Charlesworth A, Snodin C, Beer MS, Noble AJ, et al. 1992. Molecular cloning of a serotonin receptor from human brain (5‐HT1E): A fifth 5‐HT1‐like subtype. PNAS 89: 5517-5521. Meltzer HY, Li Z, Kaneda Y, Ichikawa J. 2003. Serotonin receptors: Their key role in drugs to treat schizophrenia. Prog Neuropsychopharmacol Biol Psychiat 27: 1159-1172. Messa C, Colombo C, Moresco RM, Gobbo C, Galli L, et al. 2003. 5‐HT(2A) receptor binding is reduced in drug‐naive and unchanged in SSRI‐responder depressed patients compared to healthy controls: A PET study. Psychopharmacology 167: 72-78. Mialet J, Berque‐Bestel I, Sicsic S, Langlois M, Fischmeister R, et al. 2000. Pharmacological characterization of the human 5‐HT4(d) receptor splice variant stably expressed in Chinese hamster ovary cells. Brit J Pharmacol 131: 827-835. Middlemiss DN, Fozard JR. 1983. 8‐Hydroxy‐2‐(di‐n‐propylamino)‐tetralin discriminates between subtypes of the 5‐HT1 recognition site. Eur J Pharmacol 90: 151-153. Millan MJ, Gobert A, Roux S, Porsolt R, Meneses A, et al. 2004. The serotonin1A receptor partial agonist S15535 [4‐(benzodioxan‐5‐yl)1‐(indan‐2‐yl)piperazine] enhances cholinergic transmission and cognitive function in rodents: A combined neurochemical and behavioral analysis. J Pharmacol Exp Ther 311: 190-203. Miller KJ, Teitler M. 1992. Quantitative autoradiography of 5‐ CT sensitive (5‐HT1D) and 5‐CT insensitive (5‐HT1E) serotonin receptors in human brain. Neurosci Lett 136: 223-226. Miner LAH, Backstrom JR, Sanders‐Bush E, Sesack SR. 2003. Ultrastructural localization of serotonin2A receptors in the middle layers of the rat prelimbic prefrontal cortex. Neuroscience 116: 107-117. Mitchell ES, Neumaier JF. 2005. 5‐HT6 receptors: A novel target for cognitive enhancement. Pharmacol Ther 108: 320-333. Molliver ME. 1987. Serotonergic neuronal systems: What their anatomic organization tells us about function. J Clin Psychopharmacol 7: 3S-23S. Monsma FJ, Shen Y, Ward RP, Hamblin MW, Sibley DR. 1993. Cloning and expression of a novel serotonin receptor with

8

high affinity for tricyclic psychotropic drugs. Mol Pharmacol 43: 320-327. Monti JM, Jantos H. 2006. Effects of the 5‐HT7 receptor antagonist SB‐269970 microinjected into the dorsal raphe nucleus on REM sleep in the rat. Behav Brain Res 167: 245-250. Morales M, Bloom FE. 1997. The 5‐HT3 receptor is present in different subpopulations of GABAergic neurons in the rat telencephalon. J Neurosci 17: 3157-3167. Moser PC, Bergis OE, Jegham S, Lochead A, Duconseille E, et al. 2002. SL65.0155, a novel 5‐hydroxytryptamine4 receptor partial agonist with potent cognition‐enhancing properties. J Pharmacol Exp Ther 302: 731-741. Mosher T, Hayes D, Greenshaw A. 2005. Differential effects of 5‐HT2C receptor ligands on place conditioning and locomotor activity in rats. Eur J Pharmacol 515: 107-116. Mullins UL, Gianutsos G, Eison AS. 1999. Effects of antidepressants on 5‐HT7 receptor regulation in the rat hypothalamus. Neuropsychopharmacol 21: 352-367. Nakai K, Fujii T, Fujimoto K, Suzuki T, Kawashima K. 1998. Effect of WAY‐100135 on the hippocampal acetylcholine release potentiated by 8‐OH‐DPAT, a serotonin1A receptor agonist, in normal and p‐chlorophenylalanine‐ treated rats as measured by in vivo microdialysis. Neurosci Res 31: 23-29. Nibuya M, Morinobu S, Duman RS. 1995. Regulation of BDNF and trkB mRNA in rat brain by chronic electroconvulsive seizure and antidepressant drug treatments. J Neurosci 15: 7539-7547. Niswender CM, Herrick‐Davis K, Dilley GE, Meltzer HY, Overholser JC, et al. 2001. RNA editing of the human serotonin 5‐HT2C receptor: Alterations in suicide and implications for serotonergic pharmacotherapy. Neuropsychopharmacology 24: 478-491. Noda M, Yasuda S, Okada M, Higashida H, Shimada A, et al. 2003. Recombinant human serotonin 5A receptors stably expressed in C6 glioma cells couple to multiple signal transduction pathways. J Neurochem 84: 222-232. O’Connor JJ, Kruk ZL. 1994. Effects of 21 days treatment with fluoxetine on stimulated endogenous 5‐hydroxytryptamine overflow in the rat dorsal raphe and suprachiasmatic nucleus studied using fast cyclic voltammetry in vitro. Brain Res 640: 328-335. Okubo Y, Suhara T, Suzuki K, Kobayashi K, Inoue O, et al. 2000. Serotonin 5‐HT2 receptors in schizophrenic patients studied by positron emission tomography. Life Sci 66: 2455-2464. Okuhara DY, Beck SG. 1998. Corticosteroids alter 5‐hydroxytryptamine1A receptor‐effector pathway in hippocampal subfield CA3 pyramidal cells. J Pharmacol Exp Ther 284: 1227-1233.

207

208

8

5‐Hydroxytryptamine in the central nervous system

Oliver KR, Kinsey AM, Wainwright A, Sirinathsinghji DJ. 2000. Localization of 5‐ht(5A) receptor‐like immunoreactivity in the rat brain. Brain Res 867: 131-142. Pandey G, Dwivedi Y, Rizavi HS, Ren X, Pandey SC, et al. 2002. Higher expression of serotonin 5‐HT2A receptors in the postmortem brains of teenage suicide victims. Am J Psychiat 159: 419-429. Parker RM, Bentley KR, Barnes NM. 1996. Allosteric modulation of 5‐HT3 receptors: Focus on alcohols and anaesthetic agents. Trends Pharmacol Sci 17: 95-99. Parks CL, Robinson PS, Sibille E, Shenk T, Toth M. 1998. Increased anxiety of mice lacking the serotonin1A receptor. PNAS 95: 10734-10739. Parsey RV, Hastings RS, Oquendo MA, Hu X, Goldman D, et al. 2006a. Effect of a triallelic functional polymorphism of the serotonin‐transporter‐linked promoter region on expression of serotonin transporter in the human brain. Am J Psychiatr 163: 48-51. Parsey RV, Hastings RS, Oquendo MA, Huang YY, Simpson N, et al. 2006b. Lower serotonin transporter binding potential in the human brain during major depressive episodes. Am J Psychiat 163: 52-58. Pasqualetti M, Ori M, Nardi I, Castagna M, Cassano GB, et al. 1998. Distribution of the 5‐HT5A serotonin receptor mRNA in the human brain. Mol Brain Res 56: 1-8. Passani MB, Pugliese AM, Azzurrini M, Corradetti R. 1994. Effects of DAU 6125, a novel 5‐hydroxytryptamine3 (5‐HT3) antagonist on electrophysiological properties of the rat hippocampus. Brit J Pharmacol 112: 695-703. Pazos A, Hoyer D, Palacios JM. 1984. The binding of serotonergic ligands to the porcine choroid plexus: Characterisation of a new type of serotonin recognition site. Eur J Pharmacol 106: 539-546. Pazos A, Palacios JM. 1985. Quantitative autoradiographic mapping of serotonin receptors in the rat brain. I. Serotonin‐1 receptors. Brain Res 346: 205-230. Pedigo NW, Yamamura HI, Nelson DL. 1981. Discrimination of multiple [3H]5‐hydroxytrytamine binding sites by the neuroleptics spiperone in the rat brain. J Neurochem 36: 205-230. Perez‐Garcia G, Gonzalez‐Espinoa C, Meneses A. 2006. An mRNA expression analysis of stimulation and blockade of 5‐HT7 receptors during memory consolidation. Behav Brain Res 169: 83-92. Perez‐Garcia G, Meneses A. 2005. Oral administration of the 5‐HT6 receptor antagonists SB‐357134 and SB‐399885 improves memory formation in an autoshaping learning task. Pharmacol Biochem Behav 81: 673-682. Peroutka SJ, Snyder SH. 1979. Multiple serotonin receptors: Differential binding of [3H]5‐hydroxytryptamine, [3H] lysergic acid diethylamide and [3H]spiroperidol. Mol Pharmacol 16: 687-699.

Phebus LA, Johnson KW, Zgombick JM, Gilbert PJ, Belle Van K, et al. 1997. Characterisation of LY344864 as a pharmacological tool to study 5‐HTlF receptors: Binding affinities, brain penetration and activity in the neurogenic dural inflammation model of migraine. Life Sci 61: 2117-2126. Pitchot W, Hansenne M, Pinto E, Reggers J, Fuchs S, et al. 2005. 5‐Hydroxytryptamine 1A receptors, major depression, and suicidal behavior. Biol Psychiat 58: 854-858. Pletscher A, Burkard WP, Bruderer H, Gey KF. 1963. Decrease of cerebral 5‐hydroxytryptamine and 5‐hydroxyindolacetic acid by an arylalkylamine. Life Sci 2: 828-833. Pobbe RLH, Zangrossi H. 2005. 5‐HT1A and 5‐HT2A receptors in the rat dorsal periaqueductal gray mediate the antipanic‐like effect induced by the stimulation of serotonergic neurons in the dorsal raphe nucleus. Psychopharmacology 183: 314-321. Pompeiano M, Palacios JM, Mengod G. 1994. Distribution of the serotonin 5‐HT2 receptor family mRNAs: Comparision between 5‐HT2A and 5‐HT2C receptors. Mol Brain Res 23: 163-178. Popa D, Lena C, Fabre V, Prenat C, Gingrich J, et al. 2005. Contribution of 5‐HT2 receptor subtypes to sleep–wakefulness and respiratory control, and functional adaptations in knock‐out mice lacking 5‐HT2A receptors. J Neurosci 25: 11231-11238. Porras G, Di Matteo V, De Deurwaerdere P, Esposito E, Spampinato U. 2002. Central serotonin4 receptors selectively regulate the impulse dependent exocytosis of dopamine in the rat striatum: In vivo studies with morphine, amphetamine and cocaine. Neuropharmacology 43: 1099-1109. Pratt GD, Bowery NG, Kilpatrick GJ, Leslie RA, Barnes NM, et al. 1990. Consensus meeting agrees distribution of 5‐HT3 receptors in mammalian hindbrain. Trends Pharmacol Sci 11: 135-137. Pritchett DB, Bach AWJ, Wozny M, Taleb O, Dal Toso R, et al. 1988. Structure and functional expression of a cloned rat serotonin 5HT‐2 receptor. EMBO J 7: 4135-4140. Pullagurla M, Bondareva T, Young R, Glennon RA. 2004. Modulation of the stimulus effects of (þ)amphetamine by the 5‐HT6 antagonist MS‐245. Pharmacol Biochem Behav 78: 263-268. Purselle DC, Nemeroff CB. 2003. Serotonin transporter: A potential substrate in the biology of suicide. Neuropsychopharmacology 28: 613-619. Qian Y, Melikian HE, Rye DB, Levey AI, Blakely RD. 1995. Identification and characterization of antidepressant‐sensitive serotonin transporter proteins using site‐ specific antibodies. J Neurosci 15: 1261-1274. Ramamoorthy S, Leibach FH, Mahesh VB, Ganapathy V. 1993. Partial purification and characterization of the human placental serotonin transporter. Placenta 14: 449-461.

5‐Hydroxytryptamine in the central nervous system Ramboz S, Oosting R, Amara DA, Kung HF, Blier P, et al. 1998. Serotonin receptor 1A knockout: An animal model of anxiety‐related disorder. PNAS 95: 14476-14481. Ramboz S, Saudou F, Amara DA, Belzung C, Segu L, et al. 1996. 5‐HT1B receptor knock‐out mice. Behavioral consequences. Behav Brain Res 73: 305-312. Reynolds GP, Mason SL, Meldrum A, De Keczer S, Parnes H, et al. 1995. 5‐Hydroxytryptamine (5‐HT)4 receptors in post‐mortem human brain tissue: Distribution, pharmacology and effects of neurodegenerative diseases. Brit J Pharmacol 114: 993-998. Riad M, Garcia S, Watkins KC, Jodoin N, Doucet E, Langlois X, El Mestikawy S, Hamon M, Descarries L, 2000. Somatodendritic localization of 5‐HT1A and preterminal axonal localization of 5‐HT1B serotonin receptors in adult rat brain. J Comp Neurol 417: 181-194. Rick CE, Stanford IM, Lacey MG. 1995. Excitation of rat substantia nigra pars reticulata neurons by 5‐hydroxytryptamine in vitro: Evidence for a direct action mediated by 5‐hydroxytryptamine 2C receptors. Neuroscience 69: 903-913. Riemer C, Borroni E, Levet‐Trafit B, Martin JR, Poli S, et al. 2003. Influence of the 5‐HT6 receptor on acetylcholine release in the cortex: Pharmacological characterization of 4‐(2‐bromo‐6‐pyrrolidin‐1‐ylpyridine‐4‐sulfonyl)phenylamine, a potent and selective 5‐HT6 receptor antagonist. J Med Chem 46: 1273-1276. Rios M, Lambe EK, Liu R, Teillon S, Liu J, et al. 2006. Severe deficits in 5‐HT2A‐mediated neurotransmission in BDNF conditional mutant mice. J Neurobiol 66: 408-420. Roberts AJ, Krucker T, Levy CL, Slanina KA, Sutcliffe JG, et al. 2004. Mice lacking 5‐HT7 receptors show specific impairments in contextual learning. Eur J Neurosci 19: 19131922. Roberts C, Thomas DR, Bate ST, Kew JNC. 2004. GABAergic modulation of 5‐HT7 receptor‐mediated effects on 5‐HT efflux in the guinea‐pig dorsal raphe nucleus. Neuropharmacology 46: 935-941. Rocha BA, Fumagalli F, Gainetdinov RR, Jones SR, Ator R, et al. 1998. Cocaine self‐administration in dopamine transporter knockout mice. Nat Neurosci 1: 132-137. Rousselle JC, Massot O, Fillion MP, Delepierre D, Zifa E, et al. 1996. Isolation and characterization of an endogenous peptide from rat brain interacting specifically with the 5‐HT1B receptors. J Biol Chem 271: 726-735. Routledge C, Bromidge SM, Moss SF, Price GW, Hirst W, et al. 2000. Characterization of SB‐271046: A potent, selective and orally active 5‐HT(6) receptor antagonist. Brit J Pharmacol 130: 1606-1612. Ruat M, Traifford E, Arrang JM, Tardivel‐Lacombe J, Diaz J, et al. 1993. A novel rat serotonin (5‐HT6) receptor: Molecular cloning, localization and stimulation of cAMP

8

accumulation. Biochem Biophy Res Commun 193: 268-276. Rudnick G, Clark J. 1993. From synapse to vesicle: The reuptake and storage of biogenic amine neurotransmitters. Biochim Biophys Acta 1144: 249-263. Rudnick G, Wall SC. 1992. The molecular mechanism of ‘‘ecstasy’’ [3,4‐methylenedioxymethamphetamine (MDMA)]: Serotonin transporters are targets for MDMA‐induced serotonin release. PNAS 89: 1817-1821. Santarelli L, Saxe M, Gross C, Surget A, Battaglia F, et al. 2003. Requirement of hippocampal neurogenesis for the behavioural effects of antidepressants. Science 301: 805-809. Sari Y. 2004. Serotonin1B receptors: From protein to physiological function and behaviour. Neurosci Biobehav Rev 28: 565-582. Sari Y, Lefevre K, Bancila M, Quignon M, Miquel MC, et al. 1997. Light and electron microscopic immunocytochemical visualization of 5‐HT1B receptors in the rat brain. Brain Res 760: 281-286. Sari Y, Miquel MC, Brisorgueil MJ, Ruiz G, Doucet E, et al. 1999. Cellular and subcellular localization of 5‐hydroxytryptamine1B receptors in the rat central nervous system: Immunocytochemical, autoradiographic and lesion studies. Neuroscience 88: 899-915. Sarnyai Z, Sibille EL, Pavlides C, Fenster RJ, McEwen BS, et al. 2000. Impaired hippocampal‐dependent learning and functional abnormalities in the hippocampus in mice lacking serotonin1A receptors. PNAS 97: 14731-14736. Sasaki M, Ishizaki K, Obata H, Goto F. 2001. Effects of 5‐HT2 and 5‐HT3 receptors on the modulation of nociceptive transmission in rat spinal cord according to the formalin test. Eur J Pharmacol 424: 45-52. Saudou F, Amara DA, Dierich A, Le Meur M, Ramboz S, et al. 1994. Enhanced aggressive behavior in mice lacking 5‐ HT1B receptor. Science 265: 1875-1878. Schechter LE, Smith DL, Li P, Lin Q, Rosenzweig‐Lipson S, et al. 2004. WAY‐466: Pharmacological profile of a novel and selective 5‐HT6 receptor agonist. Society for Neuroscience Meeting, 394.11. Schlicker E, Fink K, Molderings GJ, Price GW, Duckworth M, et al. 1997. Effects of selective 5‐HT1B (SB‐216 641) and 5‐HT1D (BRL 15572) receptor ligands on guinea‐pig and human 5‐HT auto‐ and heteroceptors. N‐S Arch Pharmacol 356: 321-327. Schmuck K, Ullmer C, Kalkman HO, Probst A, Lubbert H. 1996. Activation of meningeal 5‐HT2B receptors: An early step in the generation of migraine headache? Eur J Neurosci 8: 959-967. Schoeffter P, Waeber C. 1994. 5‐Hydroxytryptamine receptors with a 5‐HT6 receptor‐like profile stimulating adenylyl cyclase activity in pig caudate membranes. N‐S Arch Pharmacol 350: 356-360.

209

210

8

5‐Hydroxytryptamine in the central nervous system

Segrave R, Nathan PJ. 2005. Pindolol augmentation of selective serotonin reuptake inhibitors: Accounting for the variability of results of placebo controlled double‐blind studies in patients with major depression. Hum Psychopharmacol 20: 163-174. Sharp T, Umbers V, Hjorth S. 1996. The role of 5‐HT1A autoreceptors and alpha1 adrenoceptors in the inhibition of 5‐HT release‐II NAN‐190 and SDZ 216‐525. Neuropharmacology 35: 735-741. Sharpley AL, Elliott JM, Attenburrow MJ, Cowen PJ. 1994. Slow wave sleep in humans: Role of 5‐HT2A and 5‐HT2C receptors. Neuropharmacology 33: 467-471. Sheldon PW, Aghajanian GK. 1991. Excitatory responses to serotonin (5‐HT) in neurons of the rat piriform cortex: Evidence for mediation by 5‐HT1C receptors in pyramidal cells and 5‐HT2 receptors in interneurons. Synapse 9: 208-218. Shen RY, Andrade R. 1998. 5‐Hydroxytryptamine2 receptor facilitates GABAergic neurotransmission in rat hippocampus. J Pharmacol Exp Ther 285: 805-812. Shigematsu N, Yamamoto K, Higuchi S, Fukuda T. 2006. Novel non‐uniform distribution of serotonin transporter in the mouse hippocampus and neocortex revealed by N‐ and C‐terminal domain‐specific immunohistochemistry. Brain Res 1075: 110-116. Shinkai T, Ohmori O, Kojima H, Terao T, Suzuki T, et al. 1999. Association study of the 5‐HT6 receptor gene in schizophrenia. Am J Med Genet 88: 120-122. Shippenberg TS, Hen R, He M. 2000. Region‐specific enhancement of basal extracellular and cocaine‐evoked dopamine levels following constitutive deletion of the serotonin1B receptor. J Neurochem 75: 258-265. Sibille E, Sarnyai Z, Benjamin D, Gal J, Baker H, et al. 1997. Antisense inhibition of 5‐hydroxytryptamine2a receptor induces an antidepressant‐like effect in mice. Mol Pharmacol 52: 1056-1063. Silver H, Barash I, Aharon N, Kaplan A, Poyurovsky M. 2000. Fluvoxamine augmentation of antipsychotics improves negative symptoms in psychotic chronic schizophrenic patients: A placebo‐controlled study. Int Clin Psychopharmacol 15: 257-261. Silver H, Shmugliakov N. 1998. Augmentation with fluvoxamine but not maprotiline improves negative symptoms in treated schizophrenia. J Clin Psychopharmacol 18: 208-211. Sirota P, Mosheva T, Shabtay H, Giladi N, Korczyn AD. 2000. Use of the selective serotonin 3 receptor antagonist ondansetron in the treatment of neuroleptic‐induced tardive dyskinesia. Am J Psychiat 157: 287-289. Sleight AJ, Carolo C, Petit N, Zwingelstein C, Bourson A. 1995. Identification of 5‐hydroxytryptamine7 receptor binding sites in rat hypothalamus: Sensitivity to chronic antidepressant treatment. Mol Pharmacol 47: 99-103.

Sleight AJ, Smith RJ, Marsden CA, Palfreyman MG. 1989. The effects of chronic treatment with amitriptyline and MDL 72394 on the control of 5‐HT release in vivo. Neuropharmacology 28: 477-480. Smith BN, Sollars PJ, Dudek FE, Pickard GE. 2001. Serotonergic modulation of retinal input to the mouse suprachiasmatic nucleus mediated by 5HT1B and 5HT7 receptors. J Biol Rhythms 16: 25-38. Sollars PJ, Ogilvie MD, Simpson AM, Pickard GE. 2006. Photic entrainment is altered in the 5‐HT1B receptor knockout mouse. J Biol Rhythms 21: 21-32. Spa¨th M, Welzel D, Fa¨rber L. 2000. Treatment of chronic fatigue syndrome with 5‐HT3 receptor antagonists – preliminary results. Scand J Rheumatol 29: 72-77. Sprouse J, Reynolds L, Braselton J, Schmidt A. 2004. Serotonin‐induced phase advances of SCN neuronal firing in vitro: A possible role for 5‐HT5A receptors? Synapse 54: 111-118. Sprouse JS, Aghajanian GK. 1988. Responses of hippocampal pyramidal cells to putative serotonin 5‐HT1A and 5‐HT1B agonists: A comparative study with dorsal raphe neurons. Neuropharmacology 27: 707-715. Stanford IM, Lacey MG. 1996. Differential actions of serotonin, mediated by 5‐HT1B and 5‐HT2C receptors, on GABA‐mediated synaptic input to rat substantia nigra pars reticulata neurons in vitro. J Neurosci 16: 7566-7573. Starkey SJ, Skingle M. 1994. 5‐HT1D as well as 5‐HT1A autoreceptors modulate 5‐HT release in the guinea‐pig dorsal raphe nucleus. Neuropharmacology 33: 393-402. Stefulj J, Jernej B, Cicin‐Sain L, Rinner I, Schauenstein K. 2000. mRNA expression of serotonin receptors in cells of the immune tissues of the rat. Brain Behav Immun 14: 219-224. Stevens R, Rusch D, Solt K, Raines DE, Davies PA. 2005. Modulation of human 5‐hydroxytryptamine type 3AB receptors by volatile anesthetics and n‐alcohols. J Pharmacol Exp Ther 314: 338-345. Steward LJ, Ge J, Stowe RL, Brown DC, Bruton RK, et al. 1996. Ability of 5‐HT4 receptor ligands to modulate rat striatal dopamine release in vitro and in vivo. Brit J Pharmacol 117: 55-62. Stockmeier CA, Dilley GE, Shapiro LA, Overholser JC, Thompson PA, et al. 1997. Serotonin receptors in suicide victims with major depression. Neuropsychopharmacology 16: 162-173. Stockmeier CA, Shapiro LA, Dilley GE, Kolli TM, Friedman L, et al. 1998. Increase in serotonin‐1A autoreceptors in the midbrain of suicide victims with major depression: Postmortem evidence for decreased serotonin activity. J Neurosci 18: 7394-7401. Sur C, Betz H, Schloss P. 1996. Localization of the serotonin transporter in rat spinal cord. Eur J Neurosci 8: 2753-2757.

5‐Hydroxytryptamine in the central nervous system Suzuki M, Matsuda T, Asano S, Somboonthum P, Takuma K, et al. 1995. Increase of noradrenaline release in the hypothalamus of freely moving rat by postsynaptic 5‐hydroxytryptamine1A receptor activation. Brit J Pharmacol 115: 703-711. Suzuki T, Koyama H, Sugimoto M, Uchida I, Mashimo T. 2002. The diverse actions of volatile and gaseous anesthetics on human‐cloned 5‐hydroxytryptamine3 receptors expressed in Xenopus oocytes. Anesthesiology 96: 699-704. Svenningsson P, Chergui K, Rachleff I, Flajolet M, Zhang X, et al. 2006. Alterations in 5‐HT1B receptor function by p11 in depression‐like states. Science 311: 77-80. Tatarczynska E, Antkiewicz‐Michaluk L, Klodzinska A, Stachowicz K, Chojnacka‐Wojcik E. 2005. Antidepressant‐ like effect of the selective 5‐HT1B receptor agonist CP 94253: A possible mechanism of action. Eur J Pharmacol 516: 46-50. Tecott LH, Logue SF, Wehner JM, Kauer JA. 1998. Perturbed dentate gyrus function in serotonin 5‐HT2C receptor mutant mice. PNAS 95: 15026-15031. Tecott LH, Sun LM, Akana SF, Strack AM, Lowenstein DH, et al. 1995. Eating disorder and epilepsy in mice lacking 5HT2C serotonin receptors. Nature 374: 542-546. Terry AV, Buccafusco JL, Jackson WJ, Prendergast MA, Fontana DJ, et al. 1998. Enhanced delayed matching performance in younger and older macaques administered the 5‐HT4 receptor agonist, RS17017. Psychopharmacology 135: 407-415. The GKH, Prins J, Bleijenberg G, der Meer van JWM, 2003. The effect of granisetron, a 5‐HT3 receptor antagonist, in the treatment of chronic fatigue syndrome patients – a pilot study. Neth J Med 61: 285-289. Thomas DR. 2006. 5‐ht5A receptors as a therapeutic target. Pharmacol Ther 111: 707-714. Thomas DR, Melotto S, Massagrande M, Gribble AD, Jeffrey P, et al. 2003. SB‐656104‐A, a novel selective 5‐HT7 receptor antagonist, modulates REM sleep in rats. Brit J Pharmacol 139: 705-714. Thomas DR, Soffin EM, Roberts C, Kew JN, de la Flor RM, et al. 2006. SB‐699551‐A (3‐cyclopentyl‐N‐[2‐(dimethylamino)ethyl]‐N‐[(40 ‐{[(2‐phenylethyl)amino]methyl}‐4‐ biphenylyl)methyl]propanamide dihydrochloride), a novel 5‐ht(5A) receptor‐selective antagonist, enhances 5‐HT neuronal function: Evidence for an autoreceptor role for the 5‐ht(5A) receptor in guinea pig brain. Neuropharmacology 51: 566-577. Thorre´ K, Ebinger G, Michotte Y. 1998. 5‐HT4 receptor involvement in the serotonin‐enhanced dopamine efflux from the substantia nigra of the freely moving rat: A microdialysis study. Brain Res 796: 117-124. Tokarski K, Zahorodna A, Bobula B, Grzegorzewska M, Pitra P, et al. 2005. Repeated administration of citalopram and

8

imipramine alters the responsiveness of rat hippocampal circuitry to the activation of 5‐HT7 receptors. Eur J Pharmacol 524: 60-66. Tokarski K, Zahorodna A, Bobula B, Hess G. 2003. 5‐HT7 receptors increase the excitability of rat hippocampal CA1 pyramidal neurons. Brain Res 993: 230-234. Tomkins DM, O’Neill MF. 2000. Effect of 5‐HT(1B) receptor ligands on self‐administration of ethanol in an operant procedure in rats. Pharmacol Biochem Behav 66: 129-36. Trillat AC, Malagie´ I, Scearce K, Pons D, Anmella MC, et al. 1997. Regulation of serotonin release in the frontal cortex and ventral hippocampus of homozygous mice lacking 5‐HT lB receptors: In vivo microdialysis studies. J Neurochem 69: 2019-2025. Tsai SJ, Chiu HJ, Wang YC, Hong CJ. 1999. Association study of serotonin‐6 receptor variant (C267T) with schizophrenia and aggressive behavior. Neurosci Lett 271: 135-137. Vaidya VA, Marek GJ, Aghajanian GK, Duman RS. 1997. 5‐HT2A receptor‐mediated regulation of brain‐derived neurotrophic factor mRNA in the hippocampus and the neocortex. J Neurosci 17: 2785-2795. de Kar Van LD, Javed A, Zhang y, Serres F, Raap D, et al. 2001. 5‐HT2A receptors stimulate ACTH, corticosterone, oxytocin, renin, and prolactin release and activate hypothalamic CRF and oxytocin‐expressing cells. J Neurosci 21: 35723579. Varnas K, Hall H, Bonaventure P, Sedvall G. 2001. Autoradiographic mapping of 5‐HT(1B) and 5‐HT(1D) receptors in the post mortem human brain using [(3)H]GR 125743. Brain Res 915: 47-57. Varnas K, Halldin C, Pike VW, Hall H. 2003. Distribution of 5‐HT4 receptors in the postmortem human brain – an autoradiographic study using [125I]SB 207710. Eur Neuropsychopharmacol 13: 228-234. Varnas K, Hurd YL, Hall H. 2005. Regional expression of 5‐HT1B receptor mRNA in the human brain. Synapse 56: 21-28. Varnas K, Thomas DR, Tupala E, Tiihonen J, Hakan H. 2004. Distribution of 5‐HT7 receptors in the human brain: A preliminary autoradiographic study using [3H]SB‐ 269970. Neurosci Lett 367: 313-316. Villalobos C, Beique JC, Gingrich JA, Andrade R. 2005. Serotonergic regulation of calcium‐activated potassium currents in rodent prefrontal cortex. Eur J Neurosci 22: 1120-1126. Voigt MM, Laurie DJ, Seeburg PH, Bach A. 1991. Molecular cloning and characterization of a rat brain cDNA encoding a 5‐hydroxytryptamine1B receptor. EMBO J 10: 4017-4023. Vollenweider FX, Geyer MA. 2001. A systems model of altered consciousness: Integrating natural and drug‐induced psychoses. Brain Res Bull 56: 495-507.

211

212

8

5‐Hydroxytryptamine in the central nervous system

Waeber C, Moskowitz MA. 1995a. Autoradiographic visualisation of [3H]5‐carboxamidotryptamine binding sites in the guinea pig and rat brain. Eur J Pharmacol 283: 31-46. Waeber C, Moskowitz MA. 1995b. [3H]sumatriptan labels both 5‐HT1D and 5‐HT1F receptor binding sites in the guinea pig brain: An autoradiographic study. N‐S Arch Pharmacol 352: 263-275. Ward RP, Hamblin MW, Lachowicz JE, Hoffman BJ, Sibley DR, et al. 1995. Localization of serotonin subtype 6 receptor messenger RNA in the rat brain by in situ hybridisation histochemistry. Neuroscience 64: 1105-1111. Ward SE, Harrington FP, Gordon LJ, Hopley SC, Scott CM, et al. 2005. Discovery of the first potent, selective 5‐hydroxytryptamine1D receptor antagonist. J Med Chem 48: 3478-3480. Weber ET, Gannon RL, Rea MA. 1998. Local administration of serotonin agonists blocks light‐induced phase advances of the circadian activity rhythm in the hamster. J Biol Rhythms 13: 209-218. Welsh SE, Kachelries WJ, Romano AG, Simansky KJ, Harvey JA. 1998. Effects of LSD, ritanserin, 8‐OH‐DPAT and lisuride on classical conditioning in the rabbit. Pharmacol Biochem Behav 59: 469-475. Wilkinson LO, Middlemiss DN, Hutson PH. 1994. 5‐HT1A receptor activation increases hippocampal acetylcholine efflux and motor activity in the guinea pig: Agonist efficacy influences functional activity in vivo. J Pharmacol Exp Ther 270: 656-664. Williams GV, Rao SG, Goldman‐Rakic PS. 2002. The physiological role of 5‐HT2A receptors in working memory. J Neurosci 22: 2843-2854. Willins DL, Deutch AY, Roth BL. 1997. Serotonin 5‐HT2A receptors are expressed on pyramidal cells and interneurons in the rat cortex. Synapse 27: 79-82. Wissink S, Meijer O, Pearce D, der Burg van B, der Saag van PT. 2000. Regulation of the rat serotonin‐1A receptor gene by corticosteroids. J Biol Chem 275: 1321-1326.

Woolley ML, Bentley JC, Sleight AJ, Marsden CA, Fone KCF. 2001. A role for 5‐ht6 receptors in retention of spatial learning in the Morris water maze. Neuropharmacology 41: 210-219. Woolley ML, Marsden CA, Sleight AJ, Fone KC. 2003. Reversal of a cholinergic‐induced deficit in a rodent model of recognition memory by the selective 5‐HT6 receptor antagonist, Ro 04‐6790. Psychopharmacology 170: 358-367. Yamamoto H, Fujimiya M, Shirai Y, Nakashita M, Oyasu M, et al. 1998. Immunohistochemical localization of serotonin transporter in normal and colchicine treated rat brain. Neurosci Res 32: 305-312. Yan QS, Zheng S‐Z, Feng MJ, Yan SE. 2005. Involvement of 5‐HT1B receptors within the ventral tegmental area in ethanol‐induced increases in mesolimbic dopaminergic transmission. Brain Res 1060: 126-137. Yasuno F, Suhara T, Nakayama T, Ichimiya T, Okubo Y, et al. 2003. Inhibitory effect of hippocampal 5‐HT1A receptors on human explicit memory. Am J Psychiat 160: 334-340. Yatham LN, Liddle PF, Dennie J, Shiah IS, Adam MJ, et al. 1999. Decrease in brain serotonin 2 receptor binding in patients with major depression following desipramine treatment: A positron emission tomography study with fluorine 18‐labelled setoperone. Arch Gen Psychiat 56: 705-711. Yau JLW, Noble J, Widdowson J, Seckl JR. 1997. Impact of adrenalectomy on 5‐HT6 and 5‐HT7 receptor gene expression in the rat hippocampus. Mol Brain Res 45: 182-186. Zhao S, Edwards J, Carroll J, Wuedholz L, Millstein RA, et al. 2006. Insertion mutation at the C‐terminus of the serotonin transporter disrupts brain serotonin function and emotion‐related behaviours in mice. Neuroscience 140: 321-334.

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GABA Neurotransmission: An Overview

A. Schousboe . H. S. Waagepetersen

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GABA—from Metabolite to Neurotransmitter . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 214

2 2.1 2.2 2.3

GABA Synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 214 GAD65 and GAD67 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 214 GAD in Non‐GABAergic Neuronal Systems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 215 Glutamine as a GABA Precursor and Involvement of Mitochondria . . . . . . . . . . . . . . . . . . . . . . . . . . . 216

3 3.1 3.2

GABA Degradation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 216 GABA‐T . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 216 Oxidative Degradation: Neurons versus Astrocytes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 217

4 4.1 4.2

Inhibitors of GABA Synthesis and Degradation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 217 Carbonyl‐Trapping Agents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 217 Active Site‐Directed Inhibitors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 218

5 5.1 5.2

GABA Release . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 218 Vesicular Release . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 218 Nonvesicular Release . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 219

6 6.1 6.2

GABA Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 219 Ionotropic Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 219 Metabotropic Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 220

7 7.1 7.2 7.3 7.3.1

GABA Inactivation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 220 Receptor Desensitization and GABA Diffusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 220 High‐Affinity Plasma Membrane Transport . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 221 Inhibitors of GABA Transporters . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 221 Functional Implications of GABA Transport Inhibition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 221

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Abstract: GABA neurotransmission involves biosynthesis and metabolic degradation of GABA, its stimulus‐ coupled release and receptor interaction, as well as inactivation by high‐affinity transport systems in neuronal and astrocytic plasma membranes. These entities are summarized to provide the reader with information about the fundamental properties of these processes. List of Abbreviations: AOAA, amino-oxyacetic acid; BGT-1, betaine-GABA transporter-1; CACA, cis-4aminocrotonic acid; CAMP, cis-2-(aminomethyl)cyclopropane-1-carboxylic acid; CIT, citrate; CNS, central nervous system; EF-1502, N-[4,4-bis(3-methyl-2-thienyl)-3-butenyl]-4-(methylamino)-4,5,6,7tetrahydrobenzo[d]isoxazol-3-ol; exo-THPO, 4-amino-4,5,6,7-tetrahydro-1,2-benzo[d]isoxazol-3-ol; GABA, gamma-aminobutyric acid; GABA-T, GABA-transaminase; GAD, glutamic acid decarboxylase; GAT 1–4, GABA transporters 1–4; Glu, glutamate; OAA, oxaloacetate; PAG, phosphate activated glutaminase; SSA, succinate semialdehyde; SUC, succinate; TCA, tricarboxylic acid; THIP, 4,5,6,7-tetrahydroisoxazolo[5,4-c]pyridin-3-ol; THPO, 4,5,6,7-tetrahydroisoxazolo[4,5-c]pyridin-3-ol; 2OG, 2-oxoglutarate; 3-APPA, 3-aminopropylphosphonic acid; 3-APMPA, 3-aminopropyl(methyl)phosphinic acid

1

GABA—from Metabolite to Neurotransmitter

Gamma‐aminobutyric acid (GABA), or at physiologic pH more appropriately referred to as the zwitterion gamma‐aminobutyrate, is a plant‐associated amino acid, which in 1950 by three groups of researchers was reported to be present in brain extracts (Awapara et al., 1950; Roberts and Frankel, 1950; Udenfriend, 1950). Among these, Eugene Roberts most energetically embarked on a long‐lasting research strategy aimed at elucidating the biochemistry of this intriguing amino acid and later its association with brain function and neurotransmission. Within the next ten years, the basic biochemical pathways involving GABA had been worked out in considerable detail and a putative function in neurotransmission had been envisaged. These aspects are outlined in the proceedings of the first international conference on GABA held in 1959 (Roberts et al., 1960). The present review will briefly describe the different functional entities involved in GABAergic neurotransmission, that is, metabolic pathways (> Figure 9‐1), release processes, receptor interaction, and inactivation processes. More detailed accounts of these topics can be found in other volumes.

2

GABA Synthesis

Already, at the time of its discovery in brain tissue it was clear that GABA was produced from glutamate (Roberts and Frankel, 1950), and the synthesizing enzyme L‐glutamate decarboxylase (GAD) was subsequently extensively characterized (Roberts and Simonsen, 1963; Wu and Roberts, 1974) and purified to homogeneity (Wu et al., 1973). Polyacrylamide gel electrophoresis of the purified enzyme treated with sodium dodecyl sulphate revealed multiple bands (Matsuda et al., 1973) hinting at a possible heterogeneity of the enzyme at the molecular level.

2.1 GAD65 and GAD67 Elaborate studies of GAD regulation and subsequent cloning have demonstrated the existence of two distinct molecular forms of GAD termed GAD65 and GAD67 referring to their molecular weights of 65 kDa and 67 kDa, respectively (see Martin and Rimvall, 1993; Soghomonian and Martin, 1998). These molecular forms of GAD are encoded for by two independent genes and they exhibit different properties with regard to binding of the coenzyme pyridoxal phosphate and regulation by phosphorylation. Thus, GAD65 exists to a large extent as dormant apoenzyme, which may be rapidly activated by binding of the coenzyme whereas GAD67 mainly is found as the catalytically active holoenzyme. The two isoforms of the enzyme also exhibit differences with regard to subcellular distribution, GAD67 being cytosolic and distributed throughout the GABAergic neurons, that is, both in the cell bodies and the processes. On the contrary, GAD65 is mainly

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. Figure 9‐1 TCA cycle coupled to reactions involved in the GABA‐shunt, which constitutes an alternative pathway for the traditional TCA cycle. This involves the concerted action of the GABA‐metabolizing enzymes GAD and GABA‐T plus SSADH SSADH, succinate semialdehyde dehydrogenase; GAD, glutamic acid decarboxylase; GABA‐T, GABA‐transaminase; OAA, oxaloacetate; CIT, citrate; 2OG, 2‐oxoglutarate; SSA, succinate semialdehyde; SUC, succinate; Glu, glutamate; GABA, gamma‐aminobutyric acid

associated with the nerve endings, possibly in close association with vesicles storing GABA for neurotransmitter release. Both molecular forms are regulated by phosphorylation, GAD67 being inhibited by protein kinase A‐mediated phosphorylation and GAD65 being activated by phosphorylation. The difference in subcellular localization of the two GAD enzymes has led to speculations regarding distinctive functional roles in the biosynthesis of GABA in the cytoplasmic, metabolic pool and the neurotransmitter pool, respectively (Martin and Rimvall, 1993; Waagepetersen et al., 1999, 2001).

2.2 GAD in Non‐GABAergic Neuronal Systems Early immunohistochemical studies of the distribution of GAD in brain tissue (McLaughlin et al., 1974; Saito et al., 1974; Ribak et al., 1976) clearly associated GAD with GABAergic neuronal pathways, lending support to the general notion that GAD is a marker enzyme for GABAergic structures. Recent immunocytochemical studies have, however, provided evidence that GAD is likely to be present in certain glutamatergic neurons as well (Sloviter et al., 1996; Gutierrez, 2003). At present, the functional significance of this is poorly understood; however, it has been speculated that it may be related to the demonstration that GABA in addition to its neurotransmitter action can function as a neurodifferentiation molecule during early development of the CNS (Belhage et al., 1998; Waagepetersen et al., 1999; Fiszman and Schousboe, 2004). Interestingly, GABA mainly acts as an excitatory neurotransmitter at this early stage of development (Ben‐Ari et al., 1989; Cherubini et al., 1991; Ben‐Ari, 2002). It should also be noted that glutamatergic neurons may be able to accumulate GABA from surrounding GABAergic neurons, thereby maintaining a considerable intracellular concentration of GABA as recently demonstrated in cultures of dissociated cerebellum that consists mainly of glutamatergic neurons with a small population of GABAergic neurons (Sonnewald et al., 2004). Again, the functional significance of this can only be speculated upon.

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2.3 Glutamine as a GABA Precursor and Involvement of Mitochondria Exogenous glutamine has repeatedly been shown in brain tissue preparations to be a more efficient substrate for GABA biosynthesis than its immediate precursor, glutamate (Reubi et al., 1978; Westergaard et al., 1995). This may actually reflect the function of the GABA–glutamine–glutamate cycle in vivo in which GABA is transferred from GABAergic neurons to surrounding astrocytes where it is transaminated to succinic semialdehyde using 2‐oxoglutarate (2-OG) to generate glutamate. This is subsequently used to generate glutamine catalyzed by glutamine synthetase, and glutamine is transferred back to the GABAergic neuron where it is hydrolyzed by phosphate activated glutaminase (PAG) to form glutamate, which is decarboxylated to form GABA (Waagepetersen et al., 2003). It should be noted that these reactions do not require any anaplerotic activity since no net usage of a tricarboxylic acid (TCA) cycle constituent is needed. Hence, GABA biosynthesis may be maintained by a stoichiometric operation of the GABA–glutamine cycle and usage of acetyl CoA derived by astroglial pyruvate dehydrogenase activity. The fact that only a small fraction of the GABA neurotransmitter pool is drained from the neurons by the astrocytic uptake of GABA (Schousboe et al., 2004a) highly increases the probability that this machinery may work. The fact that PAG and GAD have different subcellular localizations in mitochondria and cytosol, respectively (Balazs et al., 1966), may impose some complications regarding the synthesis of GABA from glutamine. Although PAG is a mitochondrial enzyme it is often assumed that the product of the reaction catalyzed by PAG, glutamate, is actually liberated through the outer mitochondrial membrane since PAG may be located at the outer surface of the inner membrane (Kvamme et al., 2001). However, other evidence suggest that glutamate may be accessible for mitochondrial metabolism (Palaiologos et al., 1988; Zieminska et al., 2004). Detailed studies of labeling patterns of glutamate and GABA using [U‐13C]glucose or [U‐13C]glutamine in GABAergic neurons under conditions where GABA release occurs selectively either from the vesicular pool or the cytoplasmic pool have provided evidence that GABA synthesis particularly in the vesicular pool involves mitochondrial TCA cycle activity (Waagepetersen et al., 1999; 2001). In fact, approximately 60% of newly synthesized vesicular GABA requires TCA cycle activity. Such a biosynthetic pathway may provide GABAergic neurons with a regulatory repertoire beyond regulation of GAD activity (Waagepetersen et al., 2003).

3

GABA Degradation

As illustrated in > Figure 9‐1, GABAergic neurons are provided with enzymes allowing the TCA cycle to operate by replacing 2‐OG dehydrogenase and succinyl‐CoA synthetase steps by the concerted action of GAD, GABA‐transaminase (GABA‐T), and succinate semialdehyde dehydrogenase (SSADH). These latter reactions are called the GABA‐shunt of the TCA cycle, resulting in a small reduction of the normal ATP production from TCA cycle activity. The flux through the GABA‐shunt may account for about 10% of the activity through the complete TCA cycle (Balazs et al., 1970; Machiyama et al., 1970). It should be noted that the reactions involved in the GABA‐shunt may operate by a concerted action of GABAergic neurons and astrocytes as well as in the former cells alone.

3.1 GABA‐T The key enzyme in GABA catabolism is GABA‐T, which is localized in the mitochondrial matrix (Schousboe et al., 1977) and present in essentially all tissues including the brain (Wu et al., 1978). It has almost identical activities in astrocytes, GABAergic neurons, and glutamatergic neurons (> Table 9‐1). The enzyme was first purified to homogeneity by Schousboe et al. (1973) and characterized with regard to substrate specificity, inhibitors, and Km values for GABA and 2-OG (Schousboe et al., 1973, 1974; Bloch‐ Tardy et al., 1974; Cash et al., 1974; Maitre et al., 1975; John and Fowler, 1976). The rather low Km values for both substrates would indicate that the enzyme is functionally active in vivo, which is compatible with the fact that GABA can be metabolized to CO2 in situ (Machiyama et al., 1970; Yu and Hertz, 1983).

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. Table 9‐1 Activity of GABA‐T in mouse brain and astrocytes from mouse cerebral cortex and GABAergic and glutamatergic neurons Cell type/brain

GABA‐T activity (nmol
min1
mg1 protein)

Mouse braina GABAergic neuronsa Glutamatergic neuronsb Astrocytes (cerebral cortex)c

3.2  1.2 (7) 1.0  0.2 (4) 1.4  0.3 (5) 1.6  0.2 (7)

Note: Enzyme activity was determined in homogenates of tissue or cells using the assay for enzyme activity described by Schousboe et al. (1973). Cultures of GABAergic neurons were prepared from embryonic cerebral cortex and glutamatergic neurons from 7‐day‐old cerebellum, as described by Hertz et al. (1989a) and Schousboe et al. (1989). Astrocytes cultures were prepared from cerebral cortex of newborn mice, as described by Hertz et al. (1989b). Neuronal cultures were grown for 8–10 days and astrocytic cultures for 3 weeks before the enzyme activity was determined. Values represent averages  SEM of n experiments a Values are from Larsson et al. (1985) b Values are from A. Schousboe, unpublished

3.2 Oxidative Degradation: Neurons versus Astrocytes The demonstration of CO2 formation from [14C]GABA in neuronal preparations may be interpreted as oxidative metabolism of GABA (Yu and Hertz, 1983). As can be seen from > Figure 9‐1, production of CO2 in the TCA cycle/GABA‐shunt does not represent a net degradation of GABA, but rather a replacement of carbon atoms by acetyl CoA. Net GABA degradation would require pyruvate recycling via the action of malic enzyme (see chapter in Volume 5.5 by Sonnewald et al., 2007). It is, however, the current notion that pyruvate recycling is much less prominent in neurons than in astrocytes (Waagepetersen et al., 2003), making it unlikely that GABA may function as an important energy source in neurons. However, in astrocytes where pyruvate recycling does take place (Waagepetersen et al., 2003) GABA can be oxidatively metabolized to CO2 and water. If this happens, it is obviously not available for the operation of the GABA– glutamine cycle. Therefore, GABA, which enters the oxidative pathway via pyruvate recycling and acetyl CoA, is lost permanently from the neurotransmitter pool. This may have functional consequences for the maintenance of optimal GABAergic activity in the CNS (Schousboe et al., 2004a, b, c).

4

Inhibitors of GABA Synthesis and Degradation

The two enzymes involved in GABA metabolism, GAD and GABA‐T, belong to the family of enzymes that require pyridoxal phosphate as coenzyme. Since pyridoxal phosphate is formed from vitamin B6 by phosphorylation, a reaction catalyzed by pyridoxal‐kinase, these enzymes are generally referred to as B6‐ requiring enzymes. This common property makes both enzymes sensitive to carbonyl‐trapping agents, which form a Schiff base with the carbonyl group in pyridoxal phosphate.

4.1 Carbonyl‐Trapping Agents Hydrazine and its derivatives are highly toxic compounds, a property partly related to their ability to form hydrazones. Thus, the effects of these compounds on the GABA metabolizing enzymes have been investigated in detail (Tapia, 1975). Along this line of research it was found that amino‐oxyacetic acid (AOAA) is a highly potent inhibitor of both enzymes with a tenfold higher affinity for GABA‐T compared with GAD (Schousboe et al., 1974; Wu and Roberts, 1974).

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Hence, it is theoretically possible to use AOAA to preferentially inhibit GABA‐T but under most experimental conditions both enzymes will be affected. According to this, AOAA given to experimental animals will almost always lead to seizures, a condition associated with inhibition of GABA synthesis (Tapia and Meza‐Ruiz, 1975; Tapia et al., 1975). Another compound that was originally described as a pharmacological tool to induce seizures is mercaptopropionic acid (Sprince et al., 1969). A detailed spectroscopic and kinetic analysis of the action of 3‐mercaptopropionic acid on the purified GABA‐T from mouse brain showed that it reacts with the pyridoxal group of the coenzyme behaving as a competitive inhibitor with regard to GABA (Schousboe et al., 1974). Its Ki value for GABA‐T (13 mM) was as expected from the pharmacological action of the compound higher than that found for GAD (Wu and Roberts, 1974). Due to the pharmacological interest in compounds that might be able to effectively distinguish between GAD and GABA‐T, research programs were initiated investigating compounds that would act as active site‐ directed suicide inhibitors.

4.2 Active Site‐Directed Inhibitors From a drug development perspective, it would obviously be desirable to selectively inhibit GABA‐T since inhibition of GAD would inevitably lead to seizure activity (Tapia, 1975). Inhibition of GABA‐T could lead to increased GABA levels, thus offering protection against seizures (Tapia, 1975). Introducing alkylating entities into the GABA molecule, a substrate for GABA‐T, compounds were created, which on binding to the enzyme would form a Schiff base and simultaneously the active site would be covalently bound to the false substrate thereby irreversibly inhibiting the enzyme (Lippert et al., 1977). The most useful of this series of substrate mimetics turned out to be GABAculline and g‐vinyl GABA (Lippert et al., 1977). The latter GABA analog was subsequently developed into the first antiepileptic drug acting as a specific inhibitor of GABA‐T. These drugs lead to a significant increase in the synaptic pool of GABA (Iadarola and Gale, 1980; Wood et al., 1981; Gram et al., 1988). Interestingly, g‐vinyl GABA is more potent as an inhibitor of neuronal GABA‐T than of astroglial GABA‐T in intact cells (Larsson et al., 1986). This is not related to different affinities for GABA‐T in these cells, but may best be explained by the presence of a high‐affinity transporter for g‐vinyl GABA selectively in the neuronal plasma membrane (Schousboe et al., 1986). This transporter appears to be unrelated to the GABA transporters (Schousboe et al., 1986).

5

GABA Release

In accordance with its role as a neurotransmitter, GABA is stored in vesicles from which it can be released in a Ca2þ‐dependent manner on depolarization of the neuronal membrane (Curtis and Johnston, 1974; Schousboe et al., 1976; Otsuka, 1996). However, depending on the depolarizing conditions, GABA may also be released from the nonvesicular cytosolic pool, a mechanism that may play a physiological role (Bernath, 1991).

5.1 Vesicular Release Synaptic release of a neurotransmitter is thought to involve a vesicular pool of the transmitter and in keeping with this vesicles exist, which are equipped with transporters concentrating GABA 10‐ to 20‐fold over the cytosolic concentration (Otsuka, 1996). Vesicular, Ca2þ‐dependent GABA release has been extensively studied using a variety of brain tissue preparations such as slices, synaptosomes, and intact neurons in culture (Schousboe et al., 1976; Pin and Bockaert, 1989; Belhage et al., 1993). It may be interesting to note that in some of these preparations the relative magnitude of the vesicular release appears to be dependent on the nature of the depolarizing signal, that is, an excitatory amino acid or a high [Kþ] elicited response (Pin and Bockaert, 1989; Belhage et al., 1993).

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5.2 Nonvesicular Release As discussed in detail by Bernath (1991), nonvesicular GABA release elicited by depolarization in a non‐ Ca2þ‐dependent manner can be of considerable magnitude. This was directly demonstrated to be the case in GABAergic neurons cultured from dissociated mouse cerebral cortex. Thus, the glutamate‐induced GABA release that could be inhibited by the nontransportable GABA transport blocker N‐diphenyl–butenyl– nipecotic acid was larger than the Kþ‐stimulated, Ca2þ‐dependent release, which was not blocked by diphenyl–butenyl–nipecotic acid (Belhage et al., 1993). In addition, in other neuronal preparations, this GABA release from the nonvesicular cytosolic pool of GABA can be of a considerable magnitude. Considering the increasing evidence that nonsynaptic GABA receptors may be of significant functional importance (Mody, 2001), this GABA release, which is likely to involve primarily nonsynaptic sites, could play a prominent role in tonic GABAergic inhibition (see Sonnewald et al., 2004).

6

GABA Receptors

Although electrophysiological experiments in the 1960s had clearly demonstrated hyperpolarizing actions of GABA in neuronal preparations (Curtis and Watkins, 1960; Krnjevic and Schwartz, 1967), it took many years until experimental evidence could be provided supporting the existence of specific GABA receptors, which might mediate this hyperpolarizing action of GABA. Thus, specific binding sites representing GABA receptors were demonstrated using [3H]GABA binding by Peck et al. (1973) and by the research group of Solomon Snyder (Zukin et al., 1974; Enna and Snyder, 1975; Enna et al., 1975). A discovery that a specific binding site of the benzodiazepine tranquilizing drugs was associated with the GABA receptor (Haefely et al., 1975; Mo¨hler and Okada, 1977; Squires and Bræstrup, 1977) facilitated research related to the characterization of GABA receptors enormously. This research was further advanced by the synthesis of 4,5,6,7‐tetrahydroisoxazolo[5,4‐c]pyridin‐3‐ol (THIP) and a large number of other GABA receptor agonists and antagonists (see Krogsgaard‐Larsen et al., 2002; Frølund et al., 2004). These studies ultimately led to the cloning of a large family of GABA‐receptor subunits, the combination of which into complexes of five subunits forms functional receptors, which on agonist (GABA) binding flux Cl- through the membrane (see Jensen et al., 2005). In addition to these receptors, a second class of GABA receptors was discovered by the aid of pharmacological tools by Bowery et al. (1980). Thus, it was found that certain GABA responses could be mimicked by the lipophilic GABA analog Baclophen, an effect that could not be blocked by the classical GABA receptor antagonist bicuculline (Bowery et al., 1980). This led to the nomenclature GABAA receptors for the Cl channel‐forming receptors and GABAB for this new class of receptor that did not gate an ion channel.

6.1 Ionotropic Receptors Cloning studies have led to the identification of a total of 19 subunits that can participate in the formation of five‐membered complexes, which form an ion channel that can gate Cl ions (Olsen and Macdonald, 2002; Schousboe and Waagepetersen, 2003; Jensen et al., 2005). > Table 9‐2 delineates the subunits of the ionotropic receptors, which based on pharmacological properties are referred to as GABAA and GABAC receptors (Johnston, 1997). In addition to the agonist‐ or antagonist‐binding sites, the GABAA receptors have multiple binding sites for a variety of modulators such as benzodiazepines, b‐carbolines, barbiturates, steroids, ethanol, and Zn2þ, making the receptor a multidrug target (Johnston, 1997). The subunit composition greatly influences the pharmacological profile of the receptor leading, for example, to receptors with different sensitivity of benzodiazepines (Wafford et al., 1993a, b; McKernan and Whiting, 1996). Moreover, the affinity and the efficacy of the agonist, GABA, and its structural analogs are influenced by the subunit composition of the receptor complex (Ebert et al., 1994, 1997; Krogsgaard‐Larsen et al., 1997).

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. Table 9-2 Ionotropic GABA receptors: Subunit composition and basic pharmacology GABAA receptors

GABAC receptors Pharmacology

Pharmacology

Subunits

Agonists

Antagonist

a1–6 b1–3 g1–3 s e y p

GABA THIP Isoguvacine Isonipecotic acid Muscimol – –

Bicuculline Picrotoxinin – – – – –

Subunits

Agonists

Antagonist

r1–3

GABA CACA CAMP – – – –

3‐APMPA 3‐APPA – – – – –

– –

– – – –

Modified from Schousboe and Waagepetersen (2003) THIP, 4,5,6,7‐tetrahydroisoxazolo[5,4‐c]pyridin‐3‐ol; CACA, cis‐4‐aminocrotonic acid; CAMP, cis‐2‐(aminomethyl)cyclopropane‐1‐carboxylic acid; 3‐APMPA, 3‐aminopropyl(methyl)phosphinic acid; 3‐APPA, 3‐aminopropylphosphonic acid

This may be of functional importance for GABA‐mediated actions via synaptic and nonsynaptic receptors as these exhibit distinct differences with regard to subunit composition (Mody, 2001; Ebert et al., 2002).

6.2 Metabotropic Receptors The GABAB receptor, originally identified as a functional entity being activated by GABA and Baclofen in a bicuculline insensitive manner (Bowery et al., 1980), was shown to be coupled with G proteins and adenylate cyclase, the response of which leads to either an activation of Kþ channels with a subsequent increase in Kþ conductance and a hyperpolarization effect or a decrease in conductance of presynaptic Ca2þ channels resulting in a decreased transmitter release (Deisz, 1997). The cloning of this receptor (Klix and Bettler, 2002) has confirmed that it belongs to the 7TM superfamily of receptors and it has been shown to form a heteromeric complex of GABAB(1) and GABAB(2) subunits to be functionally active (Bettler and Bra¨uner‐Osborne, 2004). Recent knockout studies have shown that mice devoid of the GABAB(1) subunit exhibit epileptic seizures and such animals show lack of GABAB mediated responses (Prosser et al., 2001).

7

GABA Inactivation

All chemical neurotransmissions must induce a specific mechanism by which the receptor activation process can be terminated. In, by far, majority of transmitter systems this appears to be mediated by a combination of receptor desensitization and diffusion of the transmitter followed by high‐affinity transport into cellular elements lining the synaptic area.

7.1 Receptor Desensitization and GABA Diffusion GABAA receptors are characterized by a rapid desensitization, which is associated with a conformational change induced on agonist binding to the receptor complex (see Engblom et al., 2002). Since this is a rapid response it may well contribute significantly to the inactivation of the GABA‐induced hyperpolarization signal. On dissociation from the agonist binding site, GABA will move by diffusion in the narrow synaptic cleft before it reaches high‐affinity transport sites. Due to the short distances in the synaptic cleft, this process is also taking place at a rapid time scale.

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7.2 High‐Affinity Plasma Membrane Transport High‐affinity GABA transport was originally described in spinal cord, brain slices, and homogenates (Iversen and Johnston, 1971; Iversen and Bloom, 1972; Balcar et al., 1973; Beart and Johnston, 1973), and in synaptosomes and bulk‐prepared glial cells (Henn and Hamberger, 1971; Levi and Raiteri, 1973). Moreover, autoradiographic analysis of [3H]GABA uptake in brain slices have demonstrated uptake into inhibitory nerve terminals (Bloom and Iversen, 1971; Iversen and Bloom, 1972) as well as glial elements (Ho¨kfelt and Ljungdahl, 1970). Subsequent kinetic studies of [3H]GABA transport in C‐6 glioma cells and primary cultures of astrocytes confirmed the ability of astroglial cells to perform high‐affinity GABA uptake (Hutchison et al., 1974; Schrier and Thompson, 1974; Schousboe et al., 1977). Hence, it was clear that such transport systems for GABA reside in both neuronal and glial elements, a notion confirmed by immunohistochemical analysis using specific antibodies to the cloned GABA transporters (see Schousboe and Kanner, 2002; Sarup et al., 2003b; Schousboe et al., 2004a). Studies on the uptake capacity in GABAergic neurons and in astrocytes have led to the assumption that the majority of GABA released as neurotransmitter is likely to be taken up into the nerve endings while a smaller fraction will be taken up into the surrounding glial elements (Schousboe et al., 2004a). Since the first high‐affinity GABA transporter was cloned (Guastella et al., 1990) three other transporters for GABA have been cloned (see Schousboe and Kanner, 2002). In the mouse these are referred to as GAT1, GAT2, GAT3, and GAT4 (Schousboe and Kanner, 2002), where GAT2 is identical to the betaine‐ GABA transporter 1 called BGT‐1 (Liu et al., 1993). It should be noted that since this was not called GAT2 in the rat, the current nomenclature for the GABA transporters in these two species differ in the way that GAT‐2 and GAT‐3 in the rat correspond to GAT3 and GAT4, respectively, in the mouse (Schousboe and Kanner, 2002).

7.3 Inhibitors of GABA Transporters Since inhibition of GABA transporters expressed in neurons and astrocytes provides a means of pharmacological manipulation with the GABA system, much interest has been focused on pharmacological characterization of these systems (see Volume 11 in Handbook of Neurochemistry for further details). The demonstration that the GABA analogs of restricted conformation, nipecotic acid, guvacine, and 4,5,6,7‐tetrahydroisoxazolo[4,5‐c]pyridin‐3‐ol (THPO) were specific inhibitors of GABA transport without any affinity for the GABA receptors (Krogsgaard‐Larsen and Johnston, 1975) has subsequently led to the development of a large number of GABA analogs reflecting these structures (Andersen et al., 1999, 2001; Falch et al., 1999; Knutsen et al., 1999; Sarup et al., 2003a, b; Clausen et al., 2005). Though a large number of these GABA analogs reflect the structure of nipecotic acid and guvacine, an alternative avenue was taken altering the structure of THPO leading to the development of a series of analogs based on the structure of 4‐amino‐4,5,6,7‐tetrahydro‐1,2‐benzo[d]isoxazol‐3‐ol (exo‐THPO). Such analogs have been shown to have interesting pharmacological properties suggesting astrocytic GABA transport to be an important drug target (Falch et al., 1999; Sarup et al., 2003a, b; White et al., 2002, 2005; Clausen et al., 2005).

7.3.1 Functional Implications of GABA Transport Inhibition Based on the consideration that GABAergic neurotransmission to a large extent is based on reuse of GABA taken up into the presynaptic nerve ending, it was speculated that selective inhibition of astrocytic GABA uptake might provide protection against seizures (Schousboe et al., 1983). Recent pharmacological studies using GABA transport inhibitors with different affinities for neuronal and astrocytic GABA transport have provided some evidence that anticonvulsant activity of such compounds correlates much better with the ability to inhibit astrocytic GABA uptake than the ability to inhibit neuronal GABA uptake (White et al., 2002).

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It should, however, be kept in mind that Tiagabine, which is the only currently available clinically active antiepileptic drug (Kalviainen, 2002), is only marginally more potent as an inhibitor of astrocytic GABA uptake compared with neuronal uptake (Braestrup et al., 1990) and it inhibits GAT1 selectively (White et al., 2005). The recent finding that GABA analogs that inhibit GABA transport subtypes different from GAT1 (Dalby, 2003) has prompted renewed interest in this field of research leading to the discovery that inhibition of not only GAT1 but also GAT2 (BGT‐1) provides some rather interesting anticonvulsant properties. Hence, a newly synthesized exo‐THPO analog, N‐[4,4‐bis(3‐methyl‐2‐thienyl)‐3‐butenyl]‐4‐ (methylamino)‐4,5,6,7‐tetrahydrobenzo[d]isoxazol‐3‐ol (EF1502) has been shown to act synergistically with Tiagabine as an anticonvulsant, an action likely associated with inhibition of GAT2 (Clausen et al., 2005; White et al., 2005).

References Andersen KE, Lau J, Lundt BF, Petersen H, Huusfeldt PO, et al. 2001. Synthesis of novel GABA uptake inhibitors. Part 6: Preparation and evaluation of N‐Omega asymmetrically substituted nipecotic acid derivatives. Bioorg Med Chem 9: 2773-2785. Andersen KE, Sørensen JL, Huusfeldt PO, Knutsen LJ, Lau J, et al. 1999. Synthesis of novel GABA uptake inhibitors. 4. Bioisosteric transformation and successive optimization of known GABA uptake inhibitors leading to a series of potent anticonvulsant drug candidates. J Med Chem 42: 4281-4291. Awapara J, Landua AJ, Fuerst R, Seale B. 1950. Free g‐aminobutyric acid in brain. J Biol Chem 187: 35-39. Balazs R, Dahl D, Harwood JR. 1966. Subcellular distribution of enzymes of glutamate metabolism in rat brain. J Neurochem 13: 897-905. Balazs R, Machiyama Y, Hammond BJ, Julian T, Richter D. 1970. The operation of the gamma-aminobutyrate bypath of the tricarboxylic acid cycle in brain tissue in vitro. Biochem J 116: 445-461. Balcar VJ, Johnston GAR. 1973. High affinity uptake of transmitters: Studies on the uptake of L‐aspartate, GABA, L‐glutamate, and glycine in cat spinal cord. J Neurochem 20: 529-539. Beart PM, Johnston GAR. 1973. GABA uptake in rat brain slices: Inhibition by GABA analogues and by various drugs. J Neurochem 20: 319-324. Beart PM, Johnston GR. 1973. GABA uptake in rat brain slices: Inhibition by GABA analogues and by various drugs. J Neurochem 20: 319-324. Belhage B, Hansen GH, Schousboe A. 1993. Depolarization by K þ and glutamate activates different neurotransmitter release mechanisms in GABAergic neurons: Vesicular versus non‐vesicular release of GABA. Neuroscience 54: 1019-1034. Belhage B, Hansen GH, Elster E, Schousboe A. 1998. Effects of g‐aminobutyric acid (GABA) on synaptogenesis and synaptic function. Perspect Dev Neurobiol 5: 235-246.

Ben‐Ari Y. 2002. Excitatory actions of GABA during development: The nature of the nurture. Nat Rev Neurosci 3: 728-739 (Review). Ben‐Ari Y, Cherubini E, Corradetti R, Gaiarsa JL. 1989. Giant synaptic potentials in immature rat CA3 hippocampal neurons. J Physiol 416: 303-325. Bernath S. 1991. Calcium‐independent release of amino acid neurotransmitters: Fact or artifact? Prog Neurobiol 38: 57-91. Bettler B, Bra¨uner‐Osborne H. 2004. The GABAB receptor: from cloning to knockout mice. Molecular Neuropharmacology: Strategies and Methods. Schousboe A, Bra¨uner‐Osborne H, editors. Totowa, NJ: Humana Press; pp. 129-145. Bloch‐Tardy M, Rolland B, Gonnard P. 1974. Pig brain 4‐ aminobutyrate 2‐ketoglutarate transaminase. Purification, kinetics and physical properties. Biochimie 56: 823-832. Bloom FE, Iversen LL. 1971. Localizing 3H‐GABA in nerve terminals of rat cerebral cortex by electron microscopic autoradiography. Nature 229: 628-630. Bowery NG, Hill DR, Hudson AL, Doble A, Middlemiss DN, et al. 1980. (-)Baclofen decreases neurotransmitter release in the mammalian CNS by an action at a novel GABA receptor. Nature 283: 92-94. Braestrup C, Nielsen EB, Sonnewald U, Knutsen LJS, Andersen KE, et al. 1990. (R)‐N‐[4,4‐Bis(3‐methyl‐2‐ thienyl)but‐3‐en‐1‐yl]nipecotic acid binds with high affinity to the brain g‐aminobutyric acid uptake carrier. J Neurochem 54: 639-647. Cash C, Maitre M, Ciesielski L, Mandel P. 1974. Purification and partial characterisation of 4‐aminobutyrate 2‐ketoglutarae transaminase from human brain. FEBS Lett 47: 199-203. Cherubini E, Gaiarsa JL, Ben‐Ari Y. 1991. GABA: An excitatory transmitter in early postnatal life. TINS 14: 515-519. Clausen RP, Moltzen EK, Perregaard J, Lenz SM, Sanchez C, et al. 2005. Selective inhibitors of GABA uptake: Synthesis and molecular pharmacology of 3‐hydroxy‐4‐N‐methylamino‐4,5,6,7‐tetrahydro‐1,2‐benzo[d]isoxazole analogues. Bioorg Med Chem 13: 895-908.

GABA neurotransmission: An overview Curtis DR, Johnston GAR. 1974. Amino acid transmitters in the mammalian central nervous system. Ergeb Physiol 69: 97-188. Curtis DR, Watkins JC. 1960. The excitation and depression of spinal neurones by structurally related amino acids. J Neurochem 6: 117-141. Dalby NO. 2003. Inhibition of gamma‐aminobutyric acid uptake: Anatomy, physiology and effects against epileptic seizures. Eur J Pharmacol 479: 127-137. Deisz RA. 1997. Electrophysiology of GABAB receptors. The GABA Receptors, 2nd edn. Enna SJ, Bowery NG, editors. Totowa, NJ: Humana Press; pp. 157–208. Ebert B, Storustovu S, Mortensen M, Frølund B. 2002. Characterization of GABA(A) receptor ligands in the rat cortical wedge preparation: Evidence for action at extrasynaptic receptors? Br J Pharmacol 137: 1-8. Ebert B, Wafford KA, Whiting PJ, Krogsgaard‐Larsen P, Kemp JA. 1994. Molecular pharmacology of gamma‐ aminobutyric acid type A receptor agonists and partial agonists in oocytes injected with different alpha, beta, and gamma receptor subunit combinations. Mol Pharmacol 46: 957-963. Ebert B, Thompson SA, Saounatsou K, McKernan R, Krogsgaard‐Larsen P, et al. 1997. Differences in agonist/ antagonist binding affinity and receptor transduction using recombinant human gamma‐aminobutyric acid type A receptors. Mol Pharmacol 52: 1150-1156. Engblom AC, Carlson BX, Olsen RW, Schousboe A, Kristiansen U. 2002. Point mutation in the first transmembrane region of the beta2 subunit of the gamma‐aminobutyric acid type A receptor alters desensitization kinetics of gamma‐aminobutyric acid‐ and anesthetic‐induced channel gating. J Biol Chem 277: 17438-17447. Enna SJ, Kuhar MJ, Snyder SH. 1975. Regional distribution of postsynaptic receptor binding for gamma-aminobutyric acid (GABA) in monkey brain. Brain Res 93: 168-174. Enna SJ, Snyder SH. 1975. Properties of g‐aminobutyric acid (GABA) receptor binding in rat brain synaptic membrane fractions. Brain Res 100: 81-97. Falch E, Perregaard J, Frølund B, Søkilde B, Buur A, et al. 1999. Selective inhibitors of glial GABA uptake: Synthesis, absolute stereochemistry and pharmacology of the enantiomers of 3‐hydroxy‐4‐amino‐4,5,6,7‐tetrahydro‐1,2‐benzisoxazole (exo‐THPO) and analogues. J Med Chem 42: 5402-5414. Fiszman ML, Schousboe A. 2004. Role of calcium and kinases on the neurotrophic effect induced by g‐aminobutyric acid. J Neurosci Res 76: 435-441. Frølund B, Jørgensen AT, Liljefors T, Mortensen M, Krogsgaard‐Larsen P. 2004. Characterization of the GABAA receptor recognition site through ligand design

9

and pharmacophore modelling. Molecular Neuropharmacology: Strategies and Methods. Schousboe A, Bra¨uner‐Osborne H, editors. Totowa, NJ: Humana Press; pp. 113-127. Gram L, Larsson OM, Johnsen AH, Schousboe A. 1988. Effects of valproate, vigabatrin and aminooxyacetic acid on release of endogenous and exogenous GABA from cultured neurons. Epilepsy Res 2: 87-95. Guastella J, Nelson N, Nelson H, Czyzyk L, Keynan S, et al. 1990. Cloning and expression of a rat brain GABA transporter. Science 249: 1303-1306. Gutierrez R. 2003. The GABAergic phenotype of the glutamatergic granule cells of the dentate gyrus. Prog Neurobiol 71: 337-358. Haefely W, Kuskar A, Mo¨hler H, Pieri L, Polc P, et al. 1975. Possible involvement of GABA in the central actions of benzodiazepines. Adv Biochem Psychopharmacol 14: 131-152. Henn FA, Hamberger A. 1971. Glial cell function: Uptake of transmitter substances. Proc Natl Acad Sci USA 68: 2686-2690. Hertz L, Juurlink BHJ, Hertz E, Fosmark H, Schousboe A. 1989b. Preparation of primary cultures of mouse (rat) astrocytes. In: A Dissection and Tissue Culture Manual for the Nervous System. Shahar A, De Vellis J, Vernadakis A, Haber B, editors. New York: Alan R, Liss: pp. 105-108. Hertz E, Yu ACH, Hertz, L, Juurlink BHJ, Schousboe A. 1989a. Preparation of primary cultures of mouse cortical neurons. In: A Dissection and Tissue Culture Manual for the Nervous System. Shahar A, De Vellis J, Vernadakis A, Haber B, editors. New York: Alan R, Liss: 183-186. Ho¨kfelt T, Ljungdahl A˚. 1970. Cellular localization of labelled gamma‐aminobutyric acid (3H‐GABA) in rat cerebellar cortex: An autoradiographic study. Brain Res 22: 391-396. Hutchison HT, Werrbach K, Cance C, Haber B. 1974. Uptake of neurotransmitters by clonal lines of astrocytoma and neuroblastoma in culture. I. Transport of g‐aminobutyric acid. Brain Res 66: 265-274. Iadarola MJ, Gale K. 1980. Evaluation of increases in nerve terminal‐dependent vs nerve terminal‐independent compartments of GABA in vivo. Brain Res Bull 5(Suppl.)2: 13-19. Iversen LL, Bloom FE. 1972. Studies on the uptake of 3H‐ GABA and 3H‐glycine in slices and homogenates of rat brain and spinal cord by electron microscopic autoradiography. Brain Res 41: 131-143. Iversen LL, Johnston GAR. 1971. GABA uptake in rat central nervous system: Comparison of uptake in slices and homogenates and the effects of some inhibitors. J Neurochem 18: 1939-1950.

223

224

9

GABA neurotransmission: An overview

Jensen ML, Schousboe A, Ahring P. 2005. Charge selectivity of the Cys‐loop family of ligand gated ion channels. J Neurochem 92: 217-225. John RA, Fowler LJ. 1976. Kinetic and spectral properties of rabbit brain 4‐aminobutyrate aminotransferase. Biochem J 155: 645-651. Johnston GAR. 1997. Molecular biology, pharmacology, and physiology of GABAC receptors. The GABA Receptors, 2nd edn. Enna SJ, Bowery NG, editors. Totowa, NJ: Humana Press; pp. 297-323. Kalviainen R. 2002. Tiagabine. Clinical Efficacy and Use in Epilepsy. Levy RH, Mattson RH, Meldrum BS, Perucca E, editors. Philadelphia: Lippincott Williams & Wilkins; pp. 699-704. Klix N, Bettler B. 2002. Molecular structure of the GABAB receptors. Glutamate and GABA Receptors and Transporters. Egebjerg J, Schousboe A, Krogsgaard‐Larsen P, editors. London, UK: Taylor & Francis; pp. 277-286. Knutsen LJ, Andersen KE, Lau J, Lundt BF, HenryRF, et al. 1999. Synthesis of novel GABA uptake inhibitors. 3. Diaryloxime and diarylvinyl ether derivatives of nipecotic acid and guvacine as anticonvulsant agents. J Med Chem 42: 3447-3462. Krnjevic K, Schwartz S. 1967. The action of g‐aminobutyric acid on cortical neurons. Exp Brain Res 3: 320-336. Krogsgaard‐Larsen P, Johnston GAR. 1975. Inhibition of GABA uptake in rat brain slices by nipecotic acid, various isoxazoles and related compounds. J Neurochem 25: 797-802. Krogsgaard‐Larsen P, Frølund B, Ebert B. 1997. GABAA receptor agonists, partial agonists, and antagonists. The GABA Receptors, 2nd edn. Enna SJ, Bowery NG, editors. Totowa, NJ: Humana Press; pp. 37-81. Krogsgaard‐Larsen P, Frølund B, Kristiansen U, Ebert B. 2002. Ligands for the GABAA receptor complex. Glutamate and GABA Receptors and Transporters. Egebjerg J, Schousboe A, Krogsgaard‐Larsen P, editors. London, UK: Taylor & Francis; pp. 236-274. Kvamme E, Torgner IA, Roberg B. 2001. Kinetics and localization of brain phosphate activated glutaminase. J Neurosci Res 66: 951-958. Larsson OM, Drejer J, Kvamme E, Svenneby G, Hertz L, Schousboe A. 1985. Ontogenetic development of glutamate and GABA metabolizing enzymes in cultured cerebral cortex interneurons and in cerebral cortex in vivo. Int J Devl Neurosci 3: 177-185. Larsson OM, Gram L, Schousboe I, Schousboe A. 1986. Differential effect of gamma‐vinyl GABA and valproate on GABA‐transaminase from cultured neurons and astrocytes. Neuropharmacology 25: 617-625. Levi G, Raiteri M. 1973. Detectability of high and low affinity uptake systems for GABA and glutamate in rat brain slices and synaptosomes. Life Sci 12: 81-88.

Lippert B, Metcalf BW, Jung MJ, Casara P. 1977. 4‐Aminohex‐ 5‐enoic acid, a selective catalytic inhibitor of 4‐aminobutyric acid aminotransferase in mammalian brain. Eur J Biochem 74: 441-445. Liu QR, Lopez‐Corcuera B, Mandiyan S, Nelson H, Nelson N. 1993. Molecular characterization of four pharmacologically distinct g‐aminobutyric acid transporters in mouse brain. J Biol Chem 268: 2104-2112. Machiyama Y, Bala´zs R, Hammond BJ, Julian T, Richter D. 1970. The metabolism of g‐aminobutyrate and glucose in potassium ion‐stimulated brain tissue in vitro. Biochem J 116: 469-481. Maitre M, Ciesielski L, Cash C, Mandel P. 1975. Purification and studies on some properties of the 4‐aminobutyrate: 2‐ oxoglutarate transaminase from rat brain. Eur J Biochem 52: 157-169. Martin DL, Rimval K. 1993. Regulation of g‐aminobutyric acid synthesis in the brain. J Neurochem 60: 395-407. Matsuda T, Wu J‐Y, Roberts E. 1973. Electrophoresis of glutamic acid decarboxylase (EC 4.1.1.15) from mouse brain in sodium dodecyl sulphate polyacrylamide gels. J Neurochem 21: 167-172. McKernan RM, Whiting PJ. 1996. Which GABAA‐receptor subtypes really occur in the brain? Trends Neurosci 19: 139-143. McLaughlin BJ, Wood JG, Saito K, Barber R, Vaughn JE, et al. 1974. The fine structural localization of glutamate decarboxylase in synaptic terminals of rodent cerebellum. Brain Res 76: 377-391. Mo¨hler H, Okada T. 1977. Benzodiazepine receptors: Demonstration in the central nervous system. Science 198: 849-851. Mody I. 2001. Distinguishing between GABA(A) receptors responsible for tonic and phasic conductances. Neurochem Res 26: 907-913. Olsen RW, Macdonald RL. 2002. GABAA receptor complex: Structure and complex. Glutamate and GABA Receptors and Transporters. Egebjerg J, Schousboe A, Krogsgaard‐ Larsen P, editors. London, UK: Taylor & Francis; pp. 202-235. Otsuka M. 1996. Establishment of GABA as an inhibitory neurotransmitter at crustacean neuromuscular junction and in the mammalian central nervous system. GABA: Receptors, Transporters and Metabolism. Tanaka C, Bowery NG, editors. Basel: Birkha¨user Verlag; pp. 1-6. Palaiologos G, Hertz L, Schousboe A. 1988. Evidence that aspartate amino transferase activity and ketodicarboxylate carrier function are essential for biosynthesis of transmitter glutamate. J Neurochem 51: 317-320. Peck EJ, Jr, Schaeffer JM, Clark JH. 1973. g‐Aminobutyric acid, bicuculline, and post‐synaptic binding sites. Biochem Biophys Res Commun 52: 394-400.

GABA neurotransmission: An overview Pin JP, Bockaert J. 1989. Two distinct mechanisms, differentially affected by excitatory amino acids, trigger GABA release from fetal mouse striatal neurons in primary culture. J Neurosci 9: 648-656. Prosser HM, Gill CH, Hirst WD, Grau E, Robbins M, et al. 2001. Epileptogenesis and enhanced prepulse inhibition in GABAB1‐deficient mice. Mol Cell Neurosci 17: 1059-1070. Reubi JC, Van den Berg CJ, Cue´nod M. 1978. Glutamine as precursor for the GABA and glutamate transmitter pools. Neurosci Lett 10: 171-174. Ribak CE, Vaughn JE, Saito K, Barber R, Roberts E. 1976. Immunocytochemical localization of glutamate decarboxylase in rat substantia nigra. Brain Res 116: 287-298. Roberts E, Frankel S. 1950. g‐Aminobutyric acid in brain: Its formation from glutamic acid. J Biol Chem 187: 55-63. Roberts E, Simonsen DG. 1963. Some properties of L‐glutamic decarboxylase in mouse brain. Biochem Pharmacol 12: 113-134. Roberts E, Baxter CF, Van Harreveld A, Wiersma CAG, Adey WR, et al. (eds) 1960. Inhibition in the Nervous System and Gamma‐aminobutyric Acid. Oxford: Pergamon Press. Saito K, Barber R, Wu J‐Y, Matsuda T, Roberts E, et al. 1974. Immunohistochemical localization of glutamate decarboxylase in rat cerebellum. Proc Natl Acad Sci USA 71: 269-273. Sarup A, Larsson OM, Schousboe A. 2003b. GABA transporters and GABA‐transaminase as drug targets. Curr Drug Targ CNS Neurol Dis 2: 269-277. Sarup A, Larsson OM, Bolvig T, Frølund B, Krogsgaard‐ Larsen P, et al. 2003a. Effects of 3‐hydroxy‐4‐amino‐ 4,5,6,7‐tetrahydro‐1,2‐benzisoxazol (exo‐THPO) and its N‐substituted analogs on GABA transport in cultured neurons and astrocytes and by the four cloned mouse GABA transporters. Neurochem Int 43: 445-451. Schousboe A, Kanner B. 2002. GABA transporters: Functional and pharmacological properties. Glutamate and GABA Receptors and Transporters. Egebjerg J, Schousboe A, Krogsgaard‐Larsen P, editors. London, UK: Taylor & Francis Publ.; pp. 337-349. Schousboe A, Waagepetersen HS. 2003. GABA. Encyclopedia of Neuroscience. Adelman G, Smith BH, editors. New York, USA: Elsevier Science; pp. 166-176.www.neuroscion.com Schousboe A, Hertz L, Svenneby G. 1977. Uptake and metabolism of GABA in astrocytes cultured from dissociated mouse brain hemispheres. Neurochem Res 2: 217-229. Schousboe A, Larsson OM, Seiler N. 1986. Stereoselective uptake of the GABA‐transaminase inhibitors gamma‐vinyl GABA and gamma‐acetylenic GABA into neurons and astrocytes. Neurochem Res 11: 1497-1505. Schousboe A, Lisy V, Hertz L. 1976. Postnatal alterations in effects of potassium on uptake and release of glutamate and GABA in rat brain cortex slices. J Neurochem 26: 1023-1027.

9

Schousboe A, Wu J‐Y, Roberts E. 1973. Purification and characterization of the 4‐aminobutyrate‐2‐ketogluterate transaminase from mouse brain. Biochemistry 12: 2868-2873. Schousboe A, Wu J‐Y, Roberts E. 1974. Subunit structure and kinetic properties of 4‐aminobutyrate‐2‐ketoglutarate transaminase from mouse brain. J Neurochem 23: 11891195. Schousboe A, Larsson OM, Sarup A, White HS. 2004c. Role of the betaine/GABA transporter (BGT‐1/GAT2) for the control of epilepsy. Eur J Pharmacol 500: 281-287. Schousboe A, Larsson OM, Wood JD, Krogsgaard‐Larsen P. 1983. Transport and metabolism of GABA in neurons and glia: Implications for epilepsy. Epilepsia 24: 531-538. Schousboe A, Meier E, Drejer J, Hertz L. 1989. Preparation of primary cultures of mouse (rat) cerebellar granule cells. In: A Dissection and Tissue Culture Manual for the Nervous System. Shahar A, De Vellis J, Vernadakis A, Haber B, editors. New York: Alan R, Liss: pp. 203-206. Schousboe A, Sarup A, Larsson OM, White HS. 2004b. GABA transporters as drug targets for modulation of GABAergic activity. Biochem Pharmacol 68: 1557-1563. Schousboe A, Sarup A, Bak LK, Waagepetersen HS, Larsson OM. 2004a. Role of astrocytic transport processes in glutamatergic and GABAergic neurotransmission. Neurochem Int 45: 512-527. Schousboe I, Bro B, Schousboe A. 1977. Intramitochondrial localization of the 4-aminobutyrate-2-ketoglutarate transaminase from ox brain. Biochem J 162: 303-307. Schrier BK, Thompson EJ. 1974. On the role of glial cells in the mammalian nervous system. Uptake, excretion, metabolism of putative neurotransmitters by cultured glial tumor cells. J Biol Chem 249: 1769-1780. Sloviter M, Dichter MA, Rachinsky TL, Dean E, Goodman JH, et al. 1996. Basal expression and induction of glutamate decarboxylase and GABA in excitatory granule cells of the rat and monkey hippocampal dentate gyrus. J Comp Neurol 373: 593-618. Soghomonian JJ, Martin DL. 1998. Two isoforms of glutamate decarboxylase: Why? Trends Pharmacol Sci 19: 500-505. Sonnewald U, Syversen T, Schousboe A, Waagepetersen HS, Aschner M. 2007. Actions of toxins on cerebral metabolism at the cellular level. In: Handbook of Neurochemistry and Molecular Biology, Vol. 5. Lajtha A, editor. Springer-Verlag Berlin Heidelberg, pp. 569-585. Sonnewald U, Olstad E, Qu H, Babot Z, Cristo`fol R, et al. 2004. First direct demonstration of extensive GABA synthesis in mouse cerebellar neuronal cultures. J Neurochem 91: 796-803. Sprince H, Parker CM, Josephs JA, Jr, Magazino J. 1969. Convulsant activity of homocysteine and other short‐chain mercaptoacids: Protection therefrom. Ann N Y Acad Sci 166: 323-325.

225

226

9

GABA neurotransmission: An overview

Squires C, Bræstrup RF. 1977. Specific benzodiazepine receptors in rat brain characterized by high‐affinity (3H)diazepam binding. Proc Natl Acad Sci USA 74: 3805-3809. Tapia R. 1975. Biochemical pharmacology of GABA in CNS. Handbook of Psychopharmacology, Vol. 4. Iverson LL, Iverson SD, Snyder SH, editors. New York: Plunum Publ. Corp.; pp. 1-58. Tapia R, Meza‐Ruiz G. 1975. Differences in some properties of newborn and adult brain glutamate decarboxylase. J Neurobiol 6: 171-181. Tapia R, Pasantes‐Morales H, Taborda E, Pe´rez de la Mora M. 1975. Seizure susceptibility in the developing mouse and its relationship to glutamate decarboxylase in pyridoxal phosphate in brain. J Neurobiol 6: 159-170. Udenfriend S. 1950. Identification of g‐aminobutyric acid in brain by the isotope derivative method. J Biol Chem 187: 65-69. Waagepetersen HS, Sonnewald U, Schousboe A. 1999. The GABA paradox: Multiple roles as metabolite, neurotransmitter, and neurodifferentiative agent. J Neurochem 73: 1335-1342. Waagepetersen HS, Sonnewald U, Schousboe A. 2003. Compartmentation of glutamine, glutamate and GABA metabolism in neurons and astrocytes: Functional implications. Neuroscientist 9: 398-403. Waagepetersen HS, Sonnewald U, Gegelashvili G, Larsson OM, Schousboe A. 2001. Metabolic distinction between vesicular and cytosolic GABA in cultured GABAergic neurons using 13C MRS. J Neurosci Res 63: 347-355. Wafford KA, Whiting PJ, Kemp JA. 1993a. Differences in affinity and efficacy of benzodiazepine receptor ligands on recombinant GABAA receptor subtypes. Mol Pharmacol 43: 240-244. Wafford KA, Bain CJ, Whiting PJ, Kemp JA. 1993b. A functional comparison of the role of g subunits in recombinant g‐aminobutyric acid type A/benzodiazepine receptors. Mol Pharmacol 43: 437-442. Westergaard N, Sonnewald U, Petersen SB, Schousboe A. 1995. Glutamate and glutamine metabolism in cultured GABAergic neurons studied by 13C NMR spectroscopy:

Evidence for compartmentation and mitochondrial heterogeneity. Neurosci Lett 185: 24-28. White HS, Sarup A, Bolvig T, Kristensen AS, Petersen G, et al. 2002. Correlation between anticonvulsant activity and inhibitory action on glial GABA uptake of the highly selective mouse GAT1 inhibitor 3‐hydroxy‐4‐ amino‐4,5,6,7‐tetrahydro‐1,2‐benzisoxazole(exo‐THPO)and its N‐alkylated analogs. J Pharmacol Exp Ther 302: 636-644. White HS, Watson WP, Hansen S, Slough S, Sarup A, et al. 2005. First demonstration of a functional role for CNS betaine/GABA transporter (mGAT2) based on synergistic anticonvulsant action among inhibitors of mGAT1 and mGAT2. J Pharmacol Exp Ther 312: 866-874. Wood JD, Kurylo E, Tsui SK. 1981. Interactions of di‐n‐ propylacetate, gabaculine, and aminooxyacetic acid: Anticonvulsant activity and the gamma‐aminobutyrate system. J Neurochem 37: 1440-1447. Wu J‐Y, Roberts E. 1974. Properties of brain L‐glutamate decarboxylase: Inhibition studies. J Neurochem 23: 759-767. Wu J‐Y, Matsuda T, Roberts E. 1973. Purification and characterization of glutamate decarboxylase from mouse brain. J Biol Chem 248: 3039-3034. Wu J‐Y, Moss LG, Chude O. 1978. Distribution and tissue specificity of 4‐aminobutyrate‐2‐oxoglutarate aminotransferase. Neurochem Res 3: 207-219. Yu ACH, Hertz L. 1983. Metabolic sources of energy in astrocytes. Glutamine, Glutamate, and GABA in the Central Nervous System. Hertz L, Kvamme E, McGeer EG, Schousboe A, editors. New York: Alan R. Liss, Inc.; pp. 431-438. Zieminska E, Hilgier W, Waagepetersen HS, Hertz L, Sonnewald U, et al. 2004. Analysis of glutamine accumulation in rat brain mitochondria in the presence of a glutamine uptake inhibitor, histidine, reveals glutamine pools with a distinct access to deamidation. Neurochem Res 29: 2121-2123. Zukin SR, Young AB, Snyder SH. 1974. Gamma‐aminobutyric acid binding to receptor sites in the rat central nervous system. Proc Natl Acad Sci USA 71: 4802-4807.

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ATP‐Mediated Signaling in the Nervous System

B. Sperla´gh

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Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 228

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Synthesis, Utilization, and Storage of ATP in the Nervous System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 229

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The Release of ATP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 231

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The Extracellular Inactivation of ATP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 233

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ATP Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 235

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The Fast Transmitter Action of ATP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 238

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The Presynaptic Modulatory Role of ATP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 238

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The Role of ATP in Glia–Neuron and Glia–Glia Signaling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 241

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The Role of ATP in Sensory Transmission and in the Generation of Pain . . . . . . . . . . . . . . . . . . . . . 243

10 The Role of P2 Receptors in Behavioral Paradigms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 244 11 ATP as a Neuroimmunomodulator . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 244 12 Involvement of ATP Receptors CNS Diseases and their Potential Therapeutic Exploitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 245 13 Conclusions and Future Avenues . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 246

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Abstract: ATP functions as a ubiquitous signaling substance in neuronal and nonneuronal tissues. It is synthesized and stored in the nerve terminals, glial cells, and postsynaptic target cells, and is released in response to neuronal activity and a variety of other stimuli (activation of pre‐ and postsynaptic receptors, metabolic distress, inflammation, hypoosmotic stimuli, and cellular damage). ATP acts on various subtypes of ionotropic P2X and metabotropic P2Y receptors, which are widely distributed in the nervous system. P2X receptors mediate the fast transmitter action of ATP, which has been identified in a number of central and peripheral synapses. In addition, ATP modulates synaptic transmission pre‐ and postsynaptically, both in positive and negative directions via activation of P2X and P2Y receptors, respectively. Moreover, ATP acts as a transmitter not only in neuron–neuronal and neuro‐effector synapses, but also transmits signals between glial cells and neurons and within glial networks. Rapidly emerging data indicate that ATP plays an important role in sensory systems, that is, in the processing of the pain, in mechano‐ and chemosensory transduction and in the microglial response to inflammatory challenge. Finally, ATP might also act as a pathological mediator in acute and chronic neurodegeneration and in the following repair process. List of Abbreviations: Aβ, amyloid beta peptide; ABC, ATP binding cassette; ADP, adenosine 50 -diphosphate; 2-AG, 2-arachidonoylglycerol; AMP, adenosine 50 -monophosphate; AMPA, α-amino-5-hydroxy-3methyl-4-isoxazole propionic acid; Ap4A, diadenosine tetraphosphate; Ap5A, diadenosine pentaphosphate; ATP, adenosine 50 -triphosphate; CNS, central nervous system; CNT, concentrative nucleoside transporter; COX-2, cyclooxigenase-2; EEG, electroencephalography; E-NPPs, ecto-nucleotide pyrophosphatases; ENT, equilibrative nucleoside transporter; E-NTPDases, ectonucleoside triphosphate diphospho-hydrolases; ERK/JNK, extracellular signal regulated protein kinase; GABA, γ-amino-butyric acid; GPI, glycosylphosphatidyl inositol; GTP, guanosine 50 -triphosphate; HPLC, high performance liquid chromatography; IFNγ, interferon–γ; IL-1α, interleukin-1α; IL-1β, interleukin-1β; IL-6, interleukin-6; IL-18, interleukin-18; IMP, inosine monophosphate; iNOS, inducible nitric oxide synthase; LPS, bacterial lypopolisaccharide; mRNA, messenger RNA; NBMPR, nitrobenzylthioinosine; NMDA, N-methyl-D-aspartate; NO, nitric oxide; NTS, nucleus tractus solitarii; 6-OHDA, 6-hydroxidopamine; p38MAPK, p38 mitogen activated protein kinase; PNS, peripheral nervous system; PPADS, pyridoxal-phosphate-6-azophenyl-20 ,40 disulphonic acid; ROI, reactive oxygen intermediates; siRNA, small interfering RNA; TNFα, tumor necrosis factor α; UDP, uridine 50 -diphosphate; UTP, uridine 50 -triphosphate

1 Introduction ATP is well known as the universal energy currency of living cells. The first observation suggesting that ATP also plays an important signaling role, besides its central role in the cellular energy homeostasis, was reported by Drury and Szent‐Gyo¨rgyi in 1929. They observed that adenylyl compounds, including ATP, have a profound effect on heart rate and cardiovascular function (Drury and Szent-Gyo¨rgyi, 1929). This idea was rediscovered by Geoffrey Burnstock in the early 1970s, who proposed the purinergic nerve hypothesis, suggesting that ATP acts as a specific neurotransmitter in the nervous system (Burnstock, 1972). In the past three decades, this concept has gained solid experimental proof in a number of synapses of the PNS and the CNS. Moreover, it also turned out that extracellular ATP has a more versatile function in the neuronal information processing than a classical neurotransmitter, participating in pre‐ and postsynaptic neuromodulation, glia–neuron and glia–glia interactions, and neuroimmunomodulation. The purinergic signaling system also offers a number of intervention sites for therapeutic exploitation in diseases of the nervous system, which potentially could be used for drug development. There are a number of excellent review articles on different aspects of the purinergic signaling system. In this chapter, a summary is given on the current knowledge on the synthesis, storage, release, action, and inactivation of extracellular ATP in the nervous system and more detailed information can be found in specialized reviews (Sperla´gh and Vizi, 1996; Abbracchio and Burnstock, 1998; Ralevic and Burnstock, 1998; Illes et al., 2000; North and Surprenant, 2000; Sperlagh and Vizi, 2000; Zimmermann, 2000, 2001; Cunha and Ribeiro, 2000a; Cunha, 2001; Khakh, 2001; Khakh et al., 2001; Kennedy et al., 2003; Illes and Ribeiro, 2004; Kennedy, 2005; Koles et al., 2005; Franke and Illes, 2006).

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It is also important to note that although the source, release, and extracellular fate of ATP and its extracellular breakdown product, adenosine, is tightly coupled, they form separate signaling systems at the level of receptors. Therefore, the present chapter focuses only on the signaling functions mediated by ATP and other nucleotides, and those mediated by adenosine and other nucleosides will be detailed in Chapter 15.

2 Synthesis, Utilization, and Storage of ATP in the Nervous System Since ATP is ubiquitous, all metabolically active cells, including neurons, are able to synthesize ATP. The majority of ATP under normal metabolic conditions are formed from ADP by oxidative phosphorylation, in the mitochondria. In addition, ATP is also generated in a minor amount in the glycolytic pathway and in the citric acid cycle (> Figures 10-1 and > 10-2). The direct precursor of ATP formation is ADP, from which ATP is generated depending on the metabolic demand. This is the so‐called respiratory control, as the mitochondrial ATP production is coupled to the respiratory chain and is driven by the actual ADP concentration of the cell. The adenine ring of the ATP molecule is synthesized during the multiple steps of de novo purine biosynthesis from phosphoribosyl pyrophosphate, resulting in inosine monophosphate (IMP) production. IMP is then transaminated to AMP and then directly phosphorylated to ADP, serving as a substrate for mitochondrial oxidative phosphorylation (> Figure 10-1). However, the de novo purine biosynthesis is an energy‐consuming process, therefore purine salvage mechanisms also exist, which enable nerve terminals to use purines from exogenous sources. Among them, the most important is that nerve terminals take up adenosine via the nucleoside transport systems (see later) and the adenosine kinase enzyme converts adenosine to AMP, which then enters the reactions detailed earlier. Taken together, the activity of all these

. Figure 10-1 The chemical formula of adenosine 50 ‐triphosphate (ATP). The majority of ATP under normal metabolic conditions is formed from ADP, in the mitochondria by oxidative phosphorylation. In addition, ATP is also generated in a minor amount in the glycolytic pathway and in the citric acid cycle. The adenine ring of the ATP molecule is synthesized during the multiple steps of de novo purine biosynthesis from phosphoribosyl pyrophosphate, resulting in IMP production. IMP is then transaminated to AMP and then directly phosphorylated to ADP, serving as a substrate for mitochondrial oxidative phosphorylation. The ribose moiety is synthesized in the pentose phosphate pathway

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. Figure 10-2 Storage, release, and interconversion of ATP and adenosine in the synapse. Adenosine is taken up to the nerve terminal by the bidirectional, ENT transporter. Subsequently, it is rephosphorylated to AMP and ADP by adenosine kinase and adenylate kinase enzymes, respectively. The major pathway of ATP production is the mitochondrial oxidative phosphorylation, which converts ADP to ATP according to the energy demand of the cell, that is, to the actual amount of ADP. ATP is then transported out from the mitochondrion and is taken up by synaptic vesicles and released by vesicular exocytosis. In addition, ATP could be also released from the cytoplasm of pre‐ and postsynaptic cells. If ADP and AMP are accumulated in the cytoplasm, due to insufficient oxidative phosphorylation, cytosolic 50 ‐nucleotidase enzyme produces adenosine from AMP, which could leave the cell via the nucleoside carrier. ATP and ADP are hydrolyzed to AMP in the extracellular space by E‐NTPDase, E‐NPP, and by the alkaline phosphatase enzymes, and AMP is hydrolyzed to adenosine by the ecto‐50 ‐nucleotidase enzyme. In addition, there is also catalytic activity in the extracellular space for an ATP regenerating, reverse process, that is, to rephosporylate and interconvert nucleosides and nucleotides to ATP by the ecto‐ adenylate kinase enzyme. Finally, adenosine could be deaminated either intra‐ or extracellularly by adenosine deaminase enzyme giving rise to formation of inosine

reactions results in approximately 10 mM ATP concentration in the cytoplasm under normal metabolic conditions. The majority of ATP produced by the nerve terminal are used to fuel energy‐consuming cellular functions, and among them, the most important are (i) the maintenance of resting membrane potential by the Naþ/Kþ pump, (ii) the function of the other ion‐pumping mechanisms, such as the Ca2þ pumps of the plasma membrane and the mitochondria, (iii) the synthesis of neurotransmitters, receptors, ion channels, transporters, and other signaling proteins and molecules, like G proteins, protein kinases, GTP, etc., (iv) the build‐up of the vesicular proton gradient by the vacuolar Hþ ATPase, and (v) the steps of the

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exocytosis itself. Nevertheless, cytoplasmic ATP is also available for intra‐ and extracellular signaling process, and ATP is also taken up and stored in synaptic vesicles (Sperla´gh, 1996) (> Figure 10-2). ATP is the known constituent of cholinergic, noradrenergic, and serotonergic vesicles and is probably present in other types of synaptic vesicles as well. In cholinergic and noradrenergic vesicles, the amount of ATP is usually outnumbered by its cotransmitter mate. Thus, storage ratios of 3:1 to 50:1 were established, depending upon the type of vesicles, corresponding to about 1–200 mM concentration of ATP inside the vesicle. However, ATP content may differ in individual vesicles and can change under various conditions, for example, upon different patterns of neuronal activity. Although direct proof on their presence is yet to be demonstrated, it is also possible that purely ATP‐containing vesicles exist. In addition to ATP, other nucleotides are also stored in synaptic vesicles, such as ADP, AMP, UTP, Ap4A, Ap5A, and guanine nucleotides, which are thought to play role as signaling substances (Zimmermann, 2001). Although their concentration is less than that of ATP, it is still relatively high, that is, in millimolar range, and enough to serve as a pool for their release.

3 The Release of ATP The participation of ATP in the intercellular communication presumes its release to the extracellular space upon physiological and/or pathological stimuli. Indeed, a wide variety of stimuli are known to release ATP such as (i) electrical or chemical depolarization of nerve terminals, (ii) activation of cell surface receptors, (iii) mechanical stimuli, (iv) hypoxia/hypoglycemia/ischemia and the consequent cellular energy deprivation, (v) hypoosmotic challenge, (vi) inflammatory stimuli, and (vii) cellular damage. The release of ATP upon neuronal activity was demonstrated for the first time by Holton (1959) using antidromic stimulation of sensory nerves. Since then, the stimulation‐dependent release of endogenous ATP has been reported from a wide variety of in vitro brain slice and nerve terminal preparations and from isolated tissues innervated by the peripheral and autonomic nervous system, using electrical field stimulation, direct stimulation of specific neuronal pathways or chemical depolarization (cf. Sperla´gh and Vizi, 1996; Sperlagh and Vizi, 2000). The most frequently used techniques capable of detecting endogenous ATP release are the luciferin–luciferase assay and high performance liquid chromatography (HPLC) coupled with ultraviolet or fluorescent detection. The former method is preferable for rapid determination of ATP levels and is advantageous because of its high sensitivity, specificity, and simplicity. On the other hand, the HPLC method offers the opportunity to separate and identify all purine compounds released to the extracellular space. In addition, preloading the tissues with [3H]adenosine or [3H]adenine and then measuring the tritium efflux is also often used to detect [3H]purine release; however, the released radioactive label in this case is a mixture of released purines and their metabolic degradation products, and therefore it is necessary to analyze the composition of radioactive label by HPLC subsequently to identify the released purine compounds. Recently, a technical breakthrough was achieved in extracellular purine analysis by the introduction of the enzyme‐based microelectrode biosensor technique, which is able to follow ATP and adenosine release in a real‐time scale, both in vitro and in vivo. Using this method, stimulation‐dependent physiological ATP release was demonstrated during locomotor activity from spinal networks (Llaudet et al., 2005), during the hypoxic ventilatory response from the carotid body and in response to the elevation of blood pCO2 from the chemosensitive region of the medulla oblongata (Spyer et al., 2004; Gourine et al., 2005a, b), and from the retinal pigment epithelium to regulate the proliferation of the neuronal retina (Pearson et al., 2005). Another recent achievement in this area is the combination of the luciferin–luciferase assay with fluorescent imaging, whereby ATP release could be followed from cell layers or even from individual cells, for example, from Muller glial cells of the retina (Newman, 2003a). Given its ubiquitous nature, ATP could be released not only from neurons, but from any kind of cells within and outside the nervous system also. Therefore, one of the most intriguing questions is to identify the source of ATP involved in the regulation of neuronal functions. Several neuronal pathways have been identified as a source of ATP release during neuronal activity, which include the septohabenular projection, innervating the rat medial habenula (Sperlagh et al., 1995, 1997, 1998a), the ventral noradrenergic bundle, identified as a source of ATP release in the hypothalamus (Sperlagh et al., 1998b), the Shaffer collateral

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pathway providing the main excitatory input to the hippocampus (Wieraszko et al., 1989), the afferent nerve bundle projecting to the rat superior cervical ganglion (Vizi et al., 1997), and sympathetic nerves innervating the vas deferens (Kirkpatrick and Burnstock, 1987; Sperlagh and Vizi, 1992). ATP is shown to be released from these pathways in a vesicular fashion, that is, upon sodium channel activity and subsequent Ca2þ‐dependent exocytosis, alone or together with its cotransmitter mate. In other studies, no evidence was found for the neuronal origin of released ATP, which implicates that ATP may be also secreted via autocrine–paracrine pathways from nonneuronal cells (Hamann and Attwell, 1996; Juranyi et al., 1997; Sperlagh et al., 1999). Thus, besides neurons, one has to count on with glial cells, postsynaptic target cells, blood vessel endothelium, and the resident immune cells of the neural tissue, as the potential source of extracellular ATP. There are numerous stimuli that are able to release ATP from cultured glial cells, which include mechanical and inflammatory stimuli and the activation of ATP‐sensitive receptors themselves (Ballerini et al., 1996; Verderio and Matteoli, 2001; Coco et al., 2003). Nevertheless, the evidence that ATP is released upon similar stimuli from in situ glial cells is yet to be found. An additional mechanism, whereby ATP could be released in the nervous system is a receptor‐operated retrograde release from the postsynaptic target cells, which seems to be a significant mechanism in the periphery, at the neuromuscular junction (Smith, 1991; Vizi et al., 2000), and at the autonomic ganglia (Vizi et al., 1997) and neuro‐effector junctions (Vizi et al., 1992; Vizi and Sperlagh, 1999; Shinozuka et al., 2002). Postsynaptic ATP could contribute to neuronal ATP release and serves as an amplifying mechanism of the pre‐ and postsynaptic actions of extracellular ATP and its degradation product, adenosine. As ATP is a highly polarized molecule, which cannot pass freely through the cell membrane, it is also of interest to identify the mechanism, whereby it could enter the extracellular space. These include (i) vesicular exocytosis, (ii) carrier‐mediated release, (iii) release through channels and membrane pores, and (iv) cytolytic release. Vesicular exocytosis is a prototype mechanism for neurotransmitters to enter the extracellular space, which is expected to be a [Ca2þ]o‐dependent process. As ATP is the constituent of synaptic vesicles, it is reasonable to assume that exocytosis is accompanied by the release of ATP to the extracellular space. Indeed, [Ca2þ]o‐dependent ATP release in response to neuronal stimulation appears in almost all known neuro‐ neuronal and neuro‐effector connections of the nervous system (cf. Sperla´gh, 1996). Moreover, recent findings indicate that vesicular ATP release could be derived not only from nerve terminals but also from astrocytes (Coco et al., 2003). Although specific transporters, capable for the transmembrane movement of ATP, are yet to be molecularly identified in neurons, some data suggest that ATP could be also released in a carrier‐mediated manner. Especially, postsynaptic ATP release from smooth muscle cells appears to be driven by an ATP carrier, as this release, opposing from vesicular exocytosis, is strictly temperature dependent (> Figure 10-3) (Vizi and Sperlagh, 1999). In nonneuronal cells, ATP binding cassette (ABC) proteins have been implicated as an ATP transporter (al‐Awqati, 1995; Wang et al., 1996; Schwiebert, 1999), and these transporters are also expressed in glial cells (Ballerini et al., 2002) and mediate ATP release upon hypoosmotic challenge (Abdipranoto et al., 2003; Darby et al., 2003), but no evidence was found until now that these transporters are present and functional in central or peripheral nerves. Channels and pores are also potential candidates to drive the transmembrane movement of ATP. Connexin hemichannels are gap junction proteins, and their main function is to mediate electrical signaling, but they are also able to release neuroactive substances such as glutamate and ATP (Stout et al., 2004). Thus connexin hemichannels have been shown to mediate ATP release from astrocytes in response to mechanical stress (Stout et al., 2002) from xenopus oocytes (Bahima et al., 2006), from retina epithelium (Pearson et al., 2005), and from corneal endothelial cells (Gomes et al., 2005). Finally, a massive ATP release has been hypothesized to occur during any kind of cellular injury. This cytolytic ATP release may gain significance under a wide variety of pathological situations, for example, in hypoxia/ischemia, acute brain or spinal cord injury as well as in chronic neurodegenerative diseases, such as Alzheimer’s disease or sclerosis multiplex and in the generation of chronic pain, as demonstrated recently (Cook and McCleskey, 2002).

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. Figure 10-3 The a1adrenoceptor‐mediated, postsynaptic ATP release and subsequent contraction is carried out by a temperature‐dependent, carrier‐mediated mechanism. Stimulation of the postsynaptic a1‐adrenoceptors by noradrenaline (NA, 300 mM) in the guinea‐pig vas deferens elicits ATP release, measured by the luciferin– luciferase assay (a, b) and a concomitant biphasic contraction (c, d), consisting of an initial twitch and a second, tonic response with superimposed oscillations. Cooling the bath temperature from 37
C to 12
C, which prevents carrier‐mediated mechanisms, inhibits ATP release and twitch contraction, but not the second phase of the contractile response, indicating that muscle contraction itself is not carrier mediated. ATP release is expressed in pico moles per gram, whereas twitch tension is expressed in millinewton. Diagrams show the mean  SEM of 5–8 experiments, **p < 0.01, calculated by the Student’s t‐test

4 The Extracellular Inactivation of ATP ATP, if it is released to the extracellular space, is no longer stable; its action is temporally and spatially terminated by the hydrolysis by the ecto‐nucleotidases, a widely expressed series of membrane bound enzymes, capable to dephosphorylate purine nucleotides and giving rise to the formation of purine nucleosides, including adenosine.

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Several enzyme families are responsible for the extracellular degradation of ATP (> Figure 10-2). The first step of the inactivation of ATP is mediated by the family of ecto‐nucleoside triphosphate diphospho‐ hydrolases (E‐NTPDases, EC 3.6.1.5, also known as ectoATPase or apyrase) (cf. Zimmermann, 2000, 2001). E‐NTPDases have a molecular mass of 55–60 kDa in unglycosylated form, have one or two transmembrane domains, and highly conserved catalytic region faced to the extracellular space. Until now eight members of this enzyme family have been identified in molecular terms, numbered from E‐NTPDase 1 and to E‐NTPDase 8, and among them E‐NTPDase 1, 2, and 3 are the major ATP catabolizing enzyme of the brain (Kegel et al., 1997). Whereas E‐NTPDase 1 is able to hydrolyze ATP and ADP to AMP, E‐NTPDase 2 converts the nucleotide triphosphates to the respective diphosphates (cf. Zimmermann, 2000, 2001). These enzymes show widespread distribution in the brain (Wang et al., 1997; Wang and Guidotti, 1998) and have low micromolar Km for ATP and ADP giving rise to rapid and highly effective hydrolysis of ATP in almost all neuronal tissues, although Km values derived from biochemical determinations may vary between tissues and preparations. Thus, endogenous ATP is converted to adenosine to activate A1 adenosine receptors within a second in the hippocampus (Dunwiddie et al., 1997; Cunha et al., 1998) whereas the hydrolysis of ATP seems slower in other brain regions, such as the cerebral cortex (Cunha et al., 1994). Nevertheless, the short half‐life of ATP in the extracellular space could be still long enough for the activation of ionotropic ATP receptors, which act in a millisecond time scale, and probably also for metabotropic ATP receptors, which act on a slower, hundreds of millisecond–second time scale. E‐NTPDases have been shown to be upregulated after in vivo ischemia (Braun et al., 1998), indicating an increased endogenous ATP efflux to the extracellular space and the regulatory role of these enzymes between the nucleotidergic and adenosinergic signaling pathways under these conditions. In addition to the E‐NTPDase family, ATP could be also dephosphorylated by ecto‐nucleotide pyrophosphatases (E‐NPPs) and by alkaline phosphatases, both having broader substrate specificity, but also widespread tissue distribution (Zimmermann, 2000). The next step of extracellular inactivation is the hydrolysis of AMP by the ecto‐50 ‐nucleotidase (EC 3.1.3.5) enzyme, which is the rate‐limiting step giving rise to the formation of adenosine, which is a new extracellular signal, acting on its own receptors. The 50 ‐nucleotidase enzyme is a glycosylphosphatidyl inositol (GPI) anchored 62–74 kDa protein, which appears to exist mainly in homodimer form and linked to the plasma membrane with its active site exposed to the extracellular space (Zimmermann, 2001). Ecto‐ 50 ‐nucleotidase exhibits micromolar Km for AMP and is feed‐forwardly inhibited by ATP, and the synthetic analog a, b‐methylene adenosine diphosphate, which results in a delayed, burst‐like adenosine production (Cunha, 2001). It is also widely present in the brain and in the periphery, and it is predominantly associated to glial cells (Schoen et al., 1987; Grondal et al., 1988), although its expression has also been demonstrated in purified nerve terminals (Cunha et al., 1992; James and Richardson, 1993). Ecto‐50 ‐nucleotidase enzyme has been also implicated in disease conditions, for example, in epilepsy (Schoen et al., 1999; Bonan et al., 2000) and in vivo ischemia (Braun et al., 1997). In addition to enzymatic mechanisms, inactivating extracellular ATP, and giving rise to the formation of adenosine, there is also catalytic activity in the extracellular space for an ATP‐regenerating, reverse process, that is, to rephosporylate and interconvert nucleosides and nucleotides to ATP (> Figure 10-2). Although the ecto‐adenylate kinase (EC 2.7.4.3) and the ecto‐nucleoside diphosphokinase enzyme (EC 2.7.4.6), which can interconvert AMP, ADP, UDP, and UTP and ATP so far have been only demonstrated in nonneuronal cells (Harden et al., 1997; Yegutkin et al., 2002), one can assume that similar activity may also be present on the surface of neurons or glial cells. Moreover, the presence of an ectoATP:AMP phosphotransferase activity has already been verified on the surface of nerve terminals (Nagy et al., 1989; Terrian et al., 1989). Adenosine, generated by the ecto‐50 ‐nucleotidase or released on its own right, is then taken up to the nerve terminal and rapidly reincorporated to ATP stores, or deaminated extra‐ or intracellularly by the adenosine deaminase enzyme (> Figure 10-2). Specific nucleoside transporters, responsible for the uptake of adenosine, have two families: Equilibrative transporters (ENT) and concentrative transporters (CNT); the former is driven by the concentration gradient and the latter by the sodium gradient (Thorn and Jarvis, 1996; Cass et al., 1999). ENT transporters could carry different nucleosides including adenosine and inosine, but not nucleotides across the cell membrane in both directions, and are regarded as the dominant nucleoside transporters of the brain. ENT transporters seem to be widely expressed in both neuronal and

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glial elements of the CNS (Anderson et al., 1999), whereas CNT transporters appear to exhibit a more restricted localization (Anderson et al., 1996). ENT transporters have two isoforms, ENT1 and ENT2; while the former is sensitive to inhibition by the adenosine uptake inhibitor nitrobenzylthioinosine (NBMPR), the latter is not (Griffiths et al., 1997; Yao et al., 1997). On the other hand, another transporter inhibitor, dipyridamole, appears to block both ENT1 and ENT2 transporters (Lee and Jarvis, 1988). Since the intracellular adenosine level under normal metabolic state is in the micromolar range (Latini and Pedata, 2001), if the ENT transporter is loaded from the extracellular space by excess adenosine it mediates its uptake into the nerve terminals. When adenosine is taken up intracellular adenosine‐eliminating mechanisms, that is the enzymes adenosine kinase and, in a lesser amount, adenosine deaminase convert it to AMP and inosine, respectively, thereby maintaining the driving force of the carrier. On the other hand, the ENT transporter could also act in a reverse direction under certain circumstances, mediating the release of adenosine into the extracellular space (> Figure 10-2). This could occur during energy deprivation or metabolic distress, when ATP stores are depleted and AMP is generated intracellularly. Cytosolic 50 ‐nucleotidase, which has a relatively high Km for AMP (1–14 mM), becomes active under these conditions and accumulates adenosine intracellularly. The resultant intracellular adenosine accumulation then flows out to the extracellular space in a transporter‐mediated manner. Since the intracellular concentration of ATP is about 50 times higher than that of AMP, even a small level of ATP depletion causes a relatively large increase in AMP concentration and leads to subsequent adenosine efflux; therefore, this mechanism is a very sensitive sensor of intracellular metabolic distress (Cunha, 2001; Latini and Pedata, 2001). ENT transporters could also mediate homo‐ or heteroexchange, if loaded from the extracellular space with relatively high concentration of nucleotides or nucleosides. Intracellular adenosine‐eliminating mechanisms in this case are not able to keep up with the increased uptake and could reverse the transporter. ATP and other nucleotides in this way may elicit adenosine release and indirectly influence adenosine receptor‐ mediated actions (Sperlagh et al., 2003). As for extracellular deamination of adenosine, it generates inosine, which could be also taken up by the ENT transporters. The proportion of the uptake and extracellular deamination may vary between tissues; in general, deamination seems to be more important in nonneuronal tissues, for example, in the cardiovascular system.

5 ATP Receptors ATP exerts its biological action through diverse families of P2 nucleotide receptors. P2 receptors could be subdivided into two families, ionotropic (P2X) and metabotropic (P2Y) receptors (Ralevic and Burnstock, 1998). Ionotropic P2X receptors are 379–595 amino acids long ligand‐gated cation channels, having two transmembrane domains (TM1 and TM2) and a large extracellular loop (> Figure 10-4) (Valera et al., 1994; Khakh et al., 2001). The ligand‐binding domain is located within a cysteine‐rich extracellular loop between the lysine residues of 69 and 71 positions, and binding sites for different antagonists have also been identified on the extracellular domain of the receptor protein. P2X receptors are nonselective cation channels having permeability to both monovalent (Naþ, Kþ) and divalent (Ca2þ) cations. Moreover, upon prolonged or repetitive agonist application they also display the property of pore dilation which makes the channel permeable to high molecular weight cations up to 800 Da. Until now, seven individual members of this receptor family have been identified molecularly, which are numbered from P2X1 to P2X7, having individual kinetics and pharmacological phenotype (> Table 10-1) (North and Surprenant, 2000). These receptor proteins, however, do not function as individual receptors, but coassemble into various homo‐ or heterooligomeric assemblies to form functional receptors. Among possible combinations, so far 16 variations have been proved to be functional (Torres et al., 1999). These are all of the homooligomeric receptors, except P2X6, which does not function in homooligomeric form, and the rest are heterooligomers, formed from P2X1–P2X6 subunits. However, recently it has been reported that by N‐glycosylation even the homomeric P2X6 receptor can be rendered in function (Jones et al., 2004). On the other hand, the P2X7 receptor functions only in homooligomeric form and does not coassemble with other known P2X receptor subunits. Basically, P2X receptors are sensitive to ATP and ADP but not to AMP and adenosine, and the ligand‐binding profile of homomeric P2X receptors are well established (for further information, see North

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. Figure 10-4 Structure of P2X receptors. The receptor is 379–595 amino acids long, having two transmembrane domain (TM1 and TM2), intracellular N and C termini, and a large extracellular loop. The ligand‐binding domain is located within a cysteine‐rich extracellular loop between the lysine resides of 69 and 71 positions

and Surprenant, 2000). On the other hand, less is known about the pharmacology of heteromeric receptors; among them, the pharmacological profile of P2X2/3, P2X2/6, P2X1/2, P2X1/4, P2X1/5, and P2X4/6 are described (> Table 10-1) (Lewis et al., 1995; Le et al., 1998; Torres et al., 1998; King et al., 2000; Brown et al., 2002; Nicke et al., 2005). Briefly, the identification of receptor subtypes relies on their sensitivity to the agonist, a, b‐methylene ATP, as all the receptors except P2X2, P2X5, P2X7, and P2X2/6 are sensitive to this agonist, and on their sensitivity to the P2 receptor antagonists, PPADS and suramin. In addition, selective antagonists for certain subtypes of P2X receptors are also available, for example, NF449, which is a selective antagonist at the P2X1 receptor, or Brilliant Blue G, which is a potent and selective antagonist at the P2X7 receptor. As for the stoichiometry of the subunits, biochemical studies indicate that functional receptors are formed as trimers. In situ hybridization studies with specific riboprobes and immunocytochemical studies using antibodies raised against individual P2X receptor subunits revealed that all seven P2X receptors are widely expressed in the nervous system; however, the expression of individual receptor subunits are different and show region‐ and cell‐type‐specific distinct distribution (Collo et al., 1996). Among the P2X receptors, P2X2, P2X4, and P2X6 seem to be most abundantly expressed in the brain, whereas other subunits show more restricted localization (Collo et al., 1996; Vulchanova et al., 1996; Atkinson et al., 2000; Rubio and Soto, 2001). The typical localization of P2X2 receptor is on nerve terminals of the brain and the periphery (Vulchanova et al., 1997; Kanjhan et al., 1999; Atkinson et al., 2000), although it also appears postsynaptically (Rubio and Soto, 2001). P2X1 receptor has initially been suggested to be exclusively expressed on smooth muscle membrane, consistent with its role to mediate fast synaptic transmission at the autonomic neuro‐effector junction (Collo et al., 1996). However, more recent studies with more sensitive probes revealed that its expression is more widespread, that is, it is also present on the central and the peripheral neurons (Vulchanova et al., 1996; Calvert and Evans, 2004). The same holds true for P2X3 receptors, which are primarily associated to sensory pathways, but functional studies indicate that they are also expressed in other brain regions and autonomic pathways (Papp et al., 2004a; Knott et al., 2005; Rodrigues et al., 2005). P2X4 receptor shows heavy expression in several brain areas such as the cerebral cortex, the hippocampus,

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. Table 10-1 Classification of identified purine/pyrimidine receptors Purine/pyrimidine receptors Nucleotide receptors

Adenosine receptors

Ionotropic

Metabotropic

Metabotropic

Homomeric

Heteromeric

Homomeric

Heteromeric

Homomeric

P2X1 P2X2 P2X3 P2X4 P2X5 P2X6 P2X7

P2X1/P2X2 P2X2/P2X3 P2X2/P2X6 P2X4/P2X6 P2X1/P2X4 P2X1/P2X5

P2Y1 P2Y2 P2Y4 P2Y6 P2Y11 P2Y12 P2Y13 P2Y14

P2Y1/A1

A1 A2A A2B A3

Purine and pyrimidine sensitive receptors are divided into nucleotide‐ and adenosine receptors. Nucleotide receptors, which include all ATP‐sensitive receptors, can be further subdivided into subfamilies of ionotropic P2X and metabotropic P2Y receptors, whereas adenosine receptors are all metabotropic receptors. The P2X receptor family has seven individual members, which are numbered from P2X1 to P2X7. These receptor proteins, however, do not function as individual receptors, but coassemble into various homo‐ or heterooligomeric assemblies to form functional receptors. Among possible combinations, 16 variations have been proved to be functional, these are all of the homooligomeric receptors, whereas among the functional heteromeric receptors, the pharmacological phenotype of P2X2/3, P2X2/6, P2X1/2, P2X1/4, P2X1/5, and P2X4/6 are described. P2Y receptor family has eight individual members, numbered as P2Y1, P2Y2, P2Y4, P2Y6, P2Y11, P2Y12, P2Y13, and P2Y14. P2Y1 receptors could also heteromerize with A1 adenosine receptors, which results in a hybrid receptor, activated by both A1‐adenosine and P2‐receptor agonists

the thalamus, and the brainstem (Le et al., 1997) and is associated with postsynaptic specialization of synaptic contacts (Rubio and Soto, 2001). P2X5 subunits have the most restricted localization in the brain, although it shows strong representation in certain areas, for example, nucleus tractus solitarii (NTS) (Yao et al., 2001). Finally, the expression of P2X7 receptor in the nervous system is subject of a current debate, in contrast to the initial in situ hybridization studies, which proposed that P2X7 receptors are not expressed in the adult brain, except in reactive microglia and astroglia (Collo et al., 1997), immunocytochemical studies revealed a widespread presynaptic expression of P2X7 receptor immunoreactivity in a number of different brain areas, including the brainstem, the hippocampus, the cortex, the spinal cord, and the skeletal neuromuscular junction (Deuchars et al., 2001; Sperlagh et al., 2002; Atkinson et al., 2004). Although two studies demonstrated a pseudo‐immunoreactivity in the brain of P2X7 receptor knockout animals (Kukley et al., 2004; Sim et al., 2004), a more recent study indicates that the pseudo‐immunoreactivity represents a brain analog of P2X7 receptor, which shares its antibody‐binding domain with the cloned P2X7 receptor and partially retains its functionality (Sanchez‐Nogueiro et al., 2005). Whether this molecular entity is a genuine new receptor, a splice variant of P2X7 receptor, or a developmental side product, which is present only in P2X7 null mice, warrants further investigation. P2Y receptors all belong to G protein‐coupled receptors, having seven hydrophobic transmembrane domains, and possess their ATP‐binding site on the external side of TM3 and TM7 domains (von Kugelgen and Wetter, 2000; Barnard and Simon, 2001; Boarder and Webb, 2001; Communi et al., 2001; Abbracchio et al., 2003). P2Y receptor family has eight individual members, numbered as P2Y1, P2Y2, P2Y4, P2Y6, P2Y11, P2Y12, P2Y13, and P2Y14 (> Table 10-1). P2Y receptors are basically activated by adenine and uridine nucleotides, such as ATP, ADP, UDP, and UTP, but not by nucleosides and they could be further subdivided into adenine nucleotide‐preferring and uridine nucleotide‐preferring subgroups. P2Y1, P2Y11, P2Y12, and P2Y13 receptors belong to the former subgroup, whereas P2Y2, P2Y4, P2Y6, and P2Y14 receptors belong to

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the latter. Although the pharmacological phenotype of all cloned P2Y receptor subtypes are yet to be elaborated, some ligands are also available which display considerable selectivity to certain subtypes of the P2Y receptor family. Thus 2‐methylthio‐ADP is a selective agonist at the P2Y1 and P2Y12 receptors, whereas MRS2179 is a selective antagonist of P2Y1 receptors. Interestingly, P2Y1 receptors could also heteromerize with A1 adenosine receptors, which results in a hybrid receptor, activated by both A1‐adenosine and P2‐receptor agonists (> Table 10-1) (Yoshioka et al., 2001). In comparison with P2X receptors, our knowledge is less complete on the exact localization and cell type‐specific distribution of P2Y receptors. mRNA‐encoding P2Y1, P2Y12, P2Y13, and P2Y14 are undoubtedly present in the brain (Chambers et al., 2000; Communi et al., 2001; Hollopeter et al., 2001; Nicholas, 2001), and among them, P2Y1 receptor has widespread distribution also at different brain areas and associated with both neurons and astrocytes at the protein level (Moran‐Jimenez and Matute, 2000; Moore et al., 2001).

6 The Fast Transmitter Action of ATP The principal function attributed to extracellular ATP is that in activating postsynaptic P2X receptors, it acts as a fast excitatory neurotransmitter in neuro‐neuronal and neuro‐effector synapses. Thus, P2X receptor‐ gated synaptic currents were identified in numerous parts of the CNS and the PNS, and the first in this row in the brain was the identification of a P2 receptor‐mediated excitatory synaptic current in the medial habenula (Edwards et al., 1992), followed by the demonstration of similar currents in other neuro‐neuronal synapses such as the submucosal and celiac neurons (Evans et al., 1992; Silinsky et al., 1992), enteric neurons (Bardoni et al., 1997), locus coeruleus nucleus of the brainstem (Nieber et al., 1997; Poelchen et al., 2001), lateral hypothalamus (Jo and Role, 2002), rat trigeminal mesencephalic neurons (Patel et al., 2001), dorsal horn of the spinal cord (Galligan and Bertrand, 1994), CA1 and CA3 regions of the hippocampus (Pankratov et al., 1998), and the somato‐sensory cortex (Pankratov et al., 2002). Nevertheless, it should be noted that even after a decade of the first demonstration of a purinergic synapse in the brain, the number of identified purinergic currents is still relatively limited. Moreover, purinergic currents could be recorded usually only in a proportion of cells and only during evoked but not in spontaneous synaptic transmission. In general, relatively strong stimulation paradigm and simultaneous blockade of the action of other excitatory and inhibitory neurotransmitters, that is, GABA, acetylcholine, and serotonin, are also necessary conditions to observe purinergic currents (Khakh, 2001; Illes and Ribeiro, 2004). On the other hand, identification of purinergic pathways is hindered by the lack of potent and selective antagonists and the lack of specific morphological markers, which otherwise are routinely used for the identification of transmitters, for example, in case of classical transmitters or peptides. It is also unclear, whether ATP acts in these synapses as a genuine cotransmitter, released from common vesicles with its cotransmitter mate or an individual transmitter. Finally, future studies should prove the in vivo relevance of purinergic synapses. In neuro‐effector synapses, the cotransmitter role of ATP is well established, as it acts as a cotransmitter with noradrenaline at the sympathetic neuro‐effector junction, which has been demonstrated in a number of tissue preparations, including the vas deferens, several blood vessels, cutaneous microcirculation, etc. (Sneddon et al., 2000). It also acts as a cotransmitter with acetylcholine in certain parts of the parasympathetic neuro‐effector transmission sites, such as at the smooth muscle of the urinary bladder (Kennedy, 2001).

7 The Presynaptic Modulatory Role of ATP The presynaptic nerve terminal is an important checkpoint, whereby the efficacy of synaptic transmission could be locally and efficiently controlled (McGehee and Role, 1996; MacDermott et al., 1999; Vizi, 2000; Boehm and Kubista, 2002). Although originally it was suggested that ATP‐sensitive P2 receptors are located

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exclusively on postsynaptic sites, it was already recognized in the early 1990s that they are involved in the regulation of transmitter release. Thus it has been reported that a, b‐methylene ATP, an analog of ATP which is resistant to degradation, facilitates electrically evoked acetylcholine release from myenteric plexus of guinea pig, an effect which was not blocked by the antagonists of adenosine receptors, and therefore mediated by P2 receptors (Sperlagh and Vizi, 1991). Later on, this suggestion has been confirmed by electrophysiological tools (Sun and Stanley, 1996), and ATP‐activated ligand‐gated ion channels have been identified to be responsible for the facilitation acetylcholine release. Moreover, ATP is able not only to facilitate, but also directly trigger transmitter release, without preceding action potential and subsequent activation of voltage‐sensitive Ca2þ channels. Since P2X receptors have relatively high Ca2þ permeability (Rogers et al., 1997; Egan and Khakh, 2004), this property makes them capable to initiate neurotransmitter release by the Ca2þ influx through the receptor ion channel complex, provided that they are located near the release sites. Activation of P2X receptors elicits noradrenaline release from sympathetic nerve terminals of the rat (Boehm, 1999) and guinea‐pig (Sperlagh et al., 2000), as well as from central noradrenergic varicosities innervating the hippocampus (Papp et al., 2004a). As for the P2X receptor subunits involved, there is a considerable species and region heterogeneity; for example, P2X2 receptors have been suggested to be responsible for the enhancement of noradrenaline release from sympathetic nerve terminals of the rat (Boehm, 1999), whereas in the guinea pig P2X3 or the heteromeric P2X2/3 receptors have been shown to be involved (Sperlagh et al., 2000; Queiroz et al., 2003); moreover, P2X1 receptors also contribute to the response in central noradrenergic pathways (Papp et al., 2004a). In addition, P2 receptors enhance the release of serotonin from the hippocampus (Okada et al., 1999) and that of dopamine from the striatum (Zhang et al., 1995, 1996) and the nucleus accumbens (Krugel et al., 1999, 2001) in vivo, although the latter effects are thought to be mediated by P2Y receptors. The release of other transmitters, including the main excitatory and inhibitory transmitters of the brain, are also subject to facilitation by presynaptic P2X receptors both in the CNS and in the periphery, as confirmed partly by neurochemical and partly by electrophysiological methods. Activation of P2X receptors elicits glutamate release in the spinal cord (Gu and MacDermott, 1997; Nakatsuka and Gu, 2001), brainstem (Khakh and Henderson, 1998; Kato and Shigetomi, 2001; Shigetomi and Kato, 2004), and the hippocampus (Sperlagh et al., 2002; Khakh et al., 2003; Rodrigues et al., 2005; Fellin et al., 2006). As for the underlying receptor subunits involved in these effects, P2X2, P2X7 (Sperlagh et al., 2002; Khakh et al., 2003; Papp et al., 2004b; Fellin et al., 2006) (also confirmed by the use of subunit specific knockout mice), as well as P2X1, P2X3, and P2X2/3 receptors (Rodrigues et al., 2005) are all identified in the hippocampus, whereas in the spinal cord, P2X3 and an unknown receptor (Nakatsuka et al., 2003) are implicated. On the other hand, the regulation of GABA release by P2X receptors seems to be more restricted, with the exception of spinal cord (Hugel and Schlichter, 2000), cultured cortical (> Figure 10-5) (Wirkner et al., 2005) and hippocampal cells (Inoue et al., 1999), and the brainstem, where the excitatory and the inhibitory synaptic transmissions are subtype specifically facilitated, via P2X3 and P2X1 receptors, respectively (Watano et al., 2004). On the other hand, no evidence was found for a direct facilitation of GABA release by P2 receptor in the hippocampal nerve terminal preparation (Cunha and Ribeiro, 2000b). Nevertheless, the release of another inhibitory transmitter, glycine, is augmented by P2X receptor activation in the dorsal horn (Rhee et al., 2000) and in the brainstem trigeminal nucleus (Wang et al., 2001). In addition to facilitatory modulation, P2 receptors are also involved in the inhibitory modulation of the release of various transmitters and the metabotropic P2Y receptors, which are engineered to act on a longer time scale, play a major role in these actions. Hence, in the CNS, ATP inhibits the release of acetylcholine (Cunha et al., 1994), noradrenaline (von Kugelgen et al., 1994; Koch et al., 1997), serotonin (von Kugelgen et al., 1997), dopamine (Trendelenburg and Bultmann, 2000), and glutamate (Koizumi and Inoue, 1997; Inoue et al., 1999; Bennett and Boarder, 2000; Mendoza‐Fernandez et al., 2000), whereas the release of GABA seems to be not subject to inhibitory neuromodulation by P2 receptors. Convincing evidence is also available on similar modulation of acetylcholine and noradrenaline release in the periphery (for further references, see Cunha and Ribeiro, 2000a).

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. Figure 10-5 The P2X receptor agonist BzATP enhances the frequency, but not the amplitude of GABAA receptor‐mediated mIPSCs in neurons of rat cortical cell cultures (10–15 DIV) with a presynaptic site of action (from Wirkner et al., 2005, with permission from Blackwell Publishing). (a, b) Consecutive traces showing typical experiments in two individual cells. The P2X7 receptor‐selective antagonist Brilliant Blue G (BBG) alone has no effect (a), whereas it prevents the effect of BzATP on the frequency of mIPSCs (b). (c) Increase of the mean frequency (empty columns) but not amplitude (filled columns) of mIPSCs by BzATP; no increase of mIPSC amplitude in the presence of BBG is detected (n ¼ 7). The changes were expressed as percentage potentiation of the time‐ matching controls recorded in drug‐free ACSF (for control traces see a; n ¼ 5). Average values of amplitude and frequency of mIPSCs were calculated during a control period of 3 min, during the last 3 min of the subsequent application of BBG (0.3 mM) for 10 min in total, as well as during the last 3 min of further superfusion with BBG alone or in combination with BzATP (300 mM) for 10 min. *p < 0.05; statistically significant difference from zero. **p < 0.05; statistically significant difference from the effect of BzATP in the absence of BBG. (d) Effect of BzATP (300 mM) on the inward current induced by the GABAA agonist muscimol (10 mM) locally superfused for 2 s with 3‐min intervals. BzATP was superfused for 6 min during two consecutive muscimol applications. Typical recording out of five similar ones. mIPSCs were recorded at a holding potential of 60 mV. CNQX (10 mM), AP‐5 (50 mM), and TTX (0.5 mM) were all present in the medium

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8 The Role of ATP in Glia–Neuron and Glia–Glia Signaling In addition to its role as a fast transmitter and as a presynaptic modulator, rapidly emerging data indicate that ATP is an important signaling molecule in the communication between glia and neurons and within glial networks. Although glial cells are traditionally regarded as a simple support for the neuronal networks, now it has become clear that they are more active players in synaptic transmission (Araque et al., 2001; Newman, 2003b). Thus, there is a bidirectional communication between neurons and glial cells, and not only does the glia respond to signals originating from neurons, but it can also release transmitters, which then act on cell‐surface receptors present on the neuronal membrane and modulate synaptic activity pre‐ and postsynaptically. Among these gliotransmitters, ATP seems to be one of the most important, in addition to glutamate and other amino acids (> Figure 10-6) (Fields and Stevens, 2000). Hence, mechanical stimulation of astrocytes in hippocampal cell culture leads to the generation of Ca2þ waves in astrocytes, which spread by the release of ATP and subsequent activation of P2 receptors and lead to the depression of excitatory synaptic transmission between neurons (Koizumi et al., 2003). This glia‐driven synaptic depression is partly mediated by ATP itself acting on P2Y receptors and partly by adenosine acting on A1 adenosine receptors (Koizumi et al., 2003), and a similar mechanism has been demonstrated in the retina, where glial‐derived ATP depress the firing of ganglion cells via activation of A1 adenosine receptors (Newman, 2003a). Moreover, endogenous ATP, released activity dependently from hippocampal astrocytes in response to the action of glutamate on non‐NMDA receptors, seems also to act similarly causing homo‐ and heterosynaptic suppression of excitatory transmission, presumably again via its degradation product adenosine (Zhang et al., 2003). On the other hand, astrocytic ATP is also involved in the modulation of inhibitory transmission in the hippocampus, by the excitation of inhibitory interneurons via P2Y1 receptors, which leads to an increased synaptic inhibition within intact hippocampal synaptic networks (Bowser and Khakh, 2004) and in long‐term synaptic plasticity events (Pascual et al., 2005). Finally, in addition to presynaptic modulation, glial‐derived ATP could also cause enduring changes in postsynaptic efficacy; in the hypothalamic paraventricular nucleus, noradrenaline, acting on a1‐adrenoceptors, releases ATP from astrocytes, which then acting on P2X7 receptors enhance excitatory transmission postsynaptically by the activation of phosphatidylinositol 3‐kinase and subsequent insertion of AMPA receptors to the cell membrane (Gordon et al., 2005). In addition to its mediator role between glial cells and neurons, ATP is also the primary mediator of the extracellular communication between astrocytes (Guthrie et al., 1999). Astrocyte populations coordinate their functions via Ca2þ waves, and the spread of the Ca2þ signal is implemented by two ways: An intercellular pathway mediated by gap junctions and an extracellular pathway mediated by ATP and P2

. Figure 10-6 ATP is a gliotransmitter. Mechanical stimulation of astrocytes leads to the generation of Ca2þ waves in astrocytes, which spread by the release of ATP and subsequent activation of P2 receptors and lead to the depression of excitatory synaptic transmission between neurons. This glia‐driven synaptic depression is partly mediated by ATP itself acting on P2Y receptors and partly by adenosine, acting on A1 adenosine receptors. Moreover, endogenous ATP, released activity dependently from hippocampal astrocytes in response to the action of glutamate on non‐NMDA receptors, seems also to act similarly causing homo‐ and heterosynaptic suppression of excitatory transmission. On the other hand, astrocytic ATP is also involved in the modulation of inhibitory transmission, by the excitation of inhibitory interneurons via P2Y1 receptors, which leads to an increased synaptic inhibition within intact hippocampal synaptic networks. Finally, in addition to presynaptic modulation, glial‐derived ATP could also cause enduring changes in postsynaptic efficacy. In the hypothalamic paraventricular nucleus, noradrenaline, acting on a1‐adrenoceptors, releases ATP from astrocytes, which then acting on P2X7 receptors enhances excitatory transmission postsynaptically by the activation of phosphatidylinositol 3‐kinase and subsequent insertion of AMPA receptors to the cell membrane. Astrocyte‐derived ATP may also activate P2X7 receptors on microglial cells and elicit Ca2þ signals in the microglia, which eventually leads to cytolysis of this cell type. For corresponding references, see text

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. Figure 10-6 (continued)

receptors (Guthrie et al., 1999). Astrocytes communicate by calcium‐mediated signaling not only with each other but also with neighboring cells including neurons (see earlier, Koizumi et al., 2003) and microglia. Thus, astrocyte‐derived ATP activates P2X7 receptors on microglial cells and elicits Ca2þ signals in the microglia, which eventually leads to cytolysis of this cell type (Verderio and Matteoli, 2001).

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9 The Role of ATP in Sensory Transmission and in the Generation of Pain ATP and its action on ligand‐gated P2X and metabotropic P2Y receptors appear also to strongly participate in the sensory transmission, and in particular in the generation of chronic inflammatory and neuropathic pain (Chizh and Illes, 2001; Kennedy et al., 2003; Kennedy, 2005) and in the mechano‐ and chemosensory transduction (Burnstock and Wood, 1996; Burnstock, 2001, 2006). Although the algogenic action of ATP has been early recognized (Collier et al., 1966), the molecular mechanism underlying this action has only recently been understood, and several different P2 receptors seem to be involved. P2X receptors are expressed along the nociceptive pathways; mRNA encoding all subunits of the P2X receptor is expressed in sensory ganglia, including dorsal root, trigeminal, and nodose ganglia (Dunn et al., 2001; Ruan et al., 2005), and among them, the expression of mRNA‐encoding P2X3 receptor is in particular high. At the protein level, primarily, the small‐diameter IB4 expressing, capsaicin‐sensitive C neurons are those that express P2X3 receptor (Vulchanova et al., 1997; Bradbury et al., 1998), whereas functional studies indicate that P2X2, P2X2/3 (Li et al., 1999; Petruska et al., 2000), P2X1/5, and P2X4/6 receptors are expressed on the medium‐diameter capsaicin‐insensitive Ad fibers (Nakatsuka et al., 2003; Tsuzuki et al., 2003). All these P2X receptor subunits are also expressed at the central terminals of sensory afferents, and their activation releases glutamate and thereby facilitate excitatory transmission (Gu and MacDermott, 1997; Nakatsuka and Gu, 2001). As evidenced by the use of the P2X3‐selective antagonist A317491, antisense (Barclay et al., 2002; Honore et al., 2002), siRNA (Dorn et al., 2004), and knockout (Cockayne et al., 2000; Souslova et al., 2000) strategies, the activation of P2X3 receptor is involved in the generation of pain of different chronic inflammatory and neuropathic pain models, while they seem to be silent during acute pain and nociceptive reactions, which makes P2X3 receptor an attractive therapeutic target in the management of these disease states. As for the mechanism underlying the P2X3 receptor‐mediated pain, the constant activation of P2X3 receptors by ATP, released from the sensory neurons themselves (Holton, 1959) or by cytolysis (Cook and McCleskey, 2002), has been postulated, which could activate P2X3 receptors present on the nerve terminals in a self‐regenerative way and leading to the sensitization of the pathway and causing hyperalgesia and allodynia, characterized by chronic pain states (Kennedy, 2005). P2X2/3 and P2X3 receptors are also involved in the mechanosensory transduction of the bladder, where ATP is released from epithelial cells in response to bladder distension and participates in the induction of visceral pain and subsequent local reflexes (Vlaskovska et al., 2001; Burnstock, 2006; Cockayne et al., 2005). A similar role of purinergic mechanosensory transduction has also been implicated in case of a variety of other viscera, including the ureter, gut, and tooth pulp (see Burnstock, 2006). In addition to P2X3 and P2X2/3 receptors, other types of P2 receptors are also involved in sensory mechanisms and in the initiation and the regulation of pain. Hence, pharmacological blockade of P2X4 receptor reverse tactile allodynia (Tsuda et al., 2003) and genetic deletion of P2X7 receptors completely abolish chronic inflammatory and neuropathic pain in the Freund adjuvant‐induced and partial nerve ligation‐induced models, respectively (Chessell et al., 2005); however, in these latter studies, the action of ATP is attributed to P2X receptors expressed on microglial cells, which are activity dependently expressed during microglial activation and involved in the production of inflammatory mediators (see later). Finally, P2Y1 and P2Y4 receptor mRNA and protein are also heavily expressed on primary afferent nerve terminals and colocalize with P2X3 receptor (Ruan and Burnstock, 2003). However, in contrast to P2X receptors, the activation of P2Y1 receptors decreases the release of sensory transmitters, and thereby may counterbalance the algogenic action of ATP on P2X receptors (Gerevich et al., 2004). In addition to mechanosensory transduction, rapidly emerging data support a key role of extracellular ATP in central and peripheral chemosensory transduction, controlling respiration (Gourine, 2005). ATP, released from the glomus caroticum and activating P2X2 and/or P2X2/3 receptors located on the dendrites of carotid sinus nerve, controls the ventilatory response to hypoxia at the peripheral chemoreceptor area (Rong et al., 2003). Moreover, ATP is also released from the ventral surface of the central chemosensitive areas of the medulla oblongata and participates in the central chemosensory transduction evoked by the change of blood Hþ/CO2, that is, hypercapnia (Thomas et al., 1999, 2001; Thomas and Spyer, 2000;

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Gourine et al., 2005a, b), and the involvement of P2 receptors in the Hþ/CO2‐dependent regulation in the cortical excitability has also been observed (Dulla et al., 2005). Nevertheless, the identity of P2 receptors involved in the latter actions is still elusive.

10 The Role of P2 Receptors in Behavioral Paradigms Apart from data detailed earlier on sensory information processing, there is still a paucity of information on the function of ATP‐sensitive P2 receptors in complex neuronal functions such as learning, memory, and behavior. Although studies using knockout mice for one or more P2 receptor subunits have not revealed gross abnormalities in behavior (Cockayne et al., 2005), one cannot rule out the potential compensatory developmental upregulation in these studies, and this area clearly deserves further investigations, including more sensitive test batteries. Nevertheless, some hints of data indicate that P2 receptors indeed regulate various aspects of behavior. Intraaccumbal injection of the P2 receptor agonist 2‐methyl‐thio‐ATP enhances locomotor activity and elicits correspondent changes in EEG pattern, an effect abolished by 6‐OHDA pretreatment, and therefore is mediated by the mesolimbic dopaminergic pathway (Kittner et al., 2000). The P2 receptor antagonist, PPADS, prevents both the acute locomotor effects of amphetamine and the behavioral sensitization caused by repeated amphetamine injections in rats, indicating the participation of an endogenous purinergic signaling mechanism in the behavioral effect of psychostimulants (Kittner et al., 2001). Moreover, the blockade of P2 receptors by PPADS in the nucleus accumbens also suppresses feeding‐evoked dopamine release and feeding behavior (Kittner et al., 2004). On the other hand, it is also revealed that exogenous and endogenous activation of P2Y1 receptors elicits anxiety‐like behavior in the elevated plus maze test in rats through the activation of NO signaling (Kittner et al., 2003).

11 ATP as a Neuroimmunomodulator Microglial cells originate from monocyte or macrophage precursors and are regarded as the major immunocompetent cell type of the nervous system. Thus, they are rapidly activated in response to pathological signals such as ischemia and inflammation and respond with morphological changes transforming the resting ramified microglia to an amoeboid form with phagocytic activity, proliferation, and the production of a wide array of inflammatory mediators. Therefore, microglial activation is heavily implicated in the pathogenesis in CNS diseases and the following repair process. It has been known for a considerable time that microglial cells respond with both ionotropic and metabotropic response to ATP application (Walz et al., 1993; Norenberg et al., 1994). Later it was confirmed that all members of the P2 receptor family are expressed on resting and activated microglial cells kept in culture at the mRNA and/or protein level (Bianco et al., 2005; Xiang and Burnstock, 2005). Among various subtypes of the P2 receptors, the role of P2X7 receptors in microglial response is especially well delineated. Microglial cell lines respond to P2X7 receptor activation (Haas et al., 1996; Chessell et al., 1997; Visentin et al., 1999), with membrane depolarization, a sustained increase in intracellular free Ca2þ (Ferrari et al., 1996), the uptake of high molecular weight fluorescent dyes (Ferrari et al., 1996; Chessell et al., 1997), and the secretion of IL‐1b upon LPS stimulus (Ferrari et al., 1997b, c; Sanz and Di Virgilio, 2000). The central role of P2X7 receptors, as costimulators of the posttranslational processing of IL‐1b in microglial cells upon LPS challenge, has been repeatedly proven (Ferrari et al., 1997b, c; Sanz and Di Virgilio, 2000; Brough et al., 2002). The mechanism underlying ATP‐dependent IL‐1b maturation and release involves an outwardly directed Kþ conductance and the activation of the interleukin‐1‐converting enzyme (ICE, also known as caspase 1) responsible for the cleavage of pro‐IL1b to the mature, 17 kDa form (Sanz and Di Virgilio, 2000). This mechanism appears to participate not only in the exogenous but also in the endogenous activation of P2X7 receptors upon LPS challenge (Ferrari et al., 1997c). Moreover,

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after priming of the cells by ATP challenge, ADP and AMP also act as promoters of membrane currents and LPS‐induced IL‐1b secretion in these cells (Chakfe et al., 2002). In addition to IL‐1b, the synthesis and release of other cytokines and inflammatory mediators are also stimulated by P2X7 receptor activation in the microglia. Hence, ATP is a full stimulus (i.e., without the requirement of priming by LPS) to induce TNFa production via a Ca2þ‐dependent, ERK/JNK/p38 signaling pathway (Hide et al., 2000; Suzuki et al., 2004), induce cyclooxigenase‐2 (COX‐2) expression and seems to participate in the regulation of IL‐6 production (Chessell et al., 2005), although other subtypes of P2 receptors may be also involved in this latter effect (Inoue, 2002; Shigetomi and Kato, 2004). Moreover, Rampe et al. (2004) revealed that P2X7 receptors play a role in the distinct modulation of cytokine secretory pathways not only after LPS, but also upon amyloid beta peptide (Ab) preactivation. Whereas the production of IL‐1b, IL‐1a, TNFa, and IL‐18 was increased, that of IL‐6, the antiinflammatory cytokine, was attenuated under these conditions, implicating the involvement of P2X7 receptors in the pathogenesis of Alzheimer’s disease (Rampe et al., 2004). P2X7 receptor activation also induces iNOS mRNA expression and increases NO production from rat microglia (Ohtani et al., 2000), enhances IFNg‐induced iNOS expression and subsequent NO production in the murine BV‐2 microglial cell line (Gendron et al., 2003), and promotes the generation of reactive oxygen intermediates (ROI) by the p38MAPK pathway (Parvathenani et al., 2003). Finally, the activation of P2X7 receptors elicits a pronounced increase in 2‐AG secretion in the astroglial cells (Walter et al., 2004) and in the microglia (Witting et al., 2004), and low concentration of agonists stimulates the release of the neuroprotective mediator plasminogen from cultured microglia (Inoue et al., 1998). Therefore, regulation of the production of putatively protective (plasminogen, TNFa, 2‐AG) and harmful (IL‐1b, NO) inflammatory mediators by P2X7 receptors appears to follow a highly time‐ and concentration‐dependent pattern (Inoue, 2002). According to its pore‐forming property, the activation of P2X7 receptors also leads to cytolysis in an apoptotic fashion in the microglia (Ferrari et al., 1997a), which involves the proteolytic pathway of the caspase activation, although this is not an absolute requirement for the membrane damage and cytolysis (Ferrari et al., 1999; Brough et al., 2002). In addition to P2X7 receptors, other subtypes of the P2 receptor family are also involved in different aspects of neuroimmunomodulation, which responds to lower concentration of ATP (cf. McLarnon, 2005). Thus microglial Ca2þ influx could be also initiated by lower agonist concentrations, effects, which could be attributed to a non‐P2X7 ionotropic, probably P2X4 receptor activation, and there is also a metabotropic, P2Y receptor‐mediated response (Visentin et al., 1999; McLarnon, 2005). The activation of these receptors also induces COX‐2 in the human microglia (Choi et al., 2003), and P2Y1 receptors mediate the inhibition of the LPS‐induced IL‐1b, IL‐6, and TNF‐a production (Ogata et al., 2003). The level of proinflammatory cytokines, therefore seems to be differentially and oppositely regulated by ionotropic and metabotropic P2 receptors, similar to that observed in the case of the regulation of transmitter release. Finally, the microglial expression of P2X4 receptors and subsequent activation of the p38 MAP kinase pathway have been shown to be involved in the generation of inflammatory and neuropathic pain (see also Inoue et al., 2004; Guo et al., 2005; Schwab et al., 2005).

12 Involvement of ATP Receptors CNS Diseases and their Potential Therapeutic Exploitation The widespread involvement of ATP and its receptors in different neuronal functions implicate their role in the pathology of the nervous system as well. Moreover, ATP seems to play a more active pathogenetic role in a variety of CNS diseases, where the upregulation or the distinct activation of the ATP‐mediated signaling system provides a basis for disease‐selective therapeutic intervention. The disease conditions, where P2 receptors seem to play a role include ischemia or hypoxia, neurotrauma, epilepsy, Alzheimer’s and Parkinson’s disease, pain, drug addiction, and brain tumors (see for more details Koles et al., 2005; Franke and Illes, 2006).

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13 Conclusions and Future Avenues It is now clear that ATP is one of the important signaling molecules in the nervous system, which is involved in a wide variety of neuronal functions, including synaptic transmission, neuromodulation, glia–neuron interactions, and neuroimmunomodulation. Therefore, the initial idea of purinergic nervous system developed into a more diverse concept of purinergic signaling system. Nevertheless, there are a number of aspects which need further investigation. Despite the wealth of data on ATP‐mediated signaling at the molecular and the cellular levels, the present knowledge is still limited at the systems level. This holds true to any aspect of purinergic mechanism, including the release and the inactivation mechanisms and to ATP‐receptor‐mediated responses as well, which are well characterized in recombinant systems, but poorly extrapolated to in vivo conditions. The progress along this line might lead to the therapeutic utilization of purinergic signaling system, which offers a number of potential target sites for pharmacological intervention.

Acknowledgments This study was supported by grants of the Hungarian Research Foundation (OTKA T037457), Hungarian Medical Research Council (472/2003), Hungarian Research and Development Fund (NKFP1A/002/2004), and the Volkswagen Foundation (I777 854).

References Abbracchio MP, Burnstock G. 1998. Purinergic signalling: Pathophysiological roles. Jpn J Pharmacol 78: 113-145. Abbracchio MP, Boeynaems JM, Barnard EA, Boyer JL, Kennedy C, et al. 2003. Characterization of the UDP‐glucose receptor (re‐named here the P2Y14 receptor) adds diversity to the P2Y receptor family. Trends Pharmacol Sci 24: 52-55. Abdipranoto A, Liu GJ, Werry EL, Bennett MR. 2003. Mechanisms of secretion of ATP from cortical astrocytes triggered by uridine triphosphate. Neuroreport 14: 2177-2181. al‐Awqati Q. 1995. Regulation of ion channels by ABC transporters that secrete ATP [comment]. Science 269: 805-806. Anderson CM, Xiong W, Young JD, Cass CE, Parkinson FE. 1996. Demonstration of the existence of mRNAs encoding N1/cif and N2/cit sodium/nucleoside cotransporters in rat brain. Brain Res Mol Brain Res 42: 358-361. Anderson CM, Xiong W, Geiger JD, Young JD, Cass CE, et al. 1999. Distribution of equilibrative, nitrobenzylthioinosine‐ sensitive nucleoside transporters (ENT1) in brain. J Neurochem 73: 867-873. Araque A, Carmignoto G, Haydon PG. 2001. Dynamic signaling between astrocytes and neurons. Annu Rev Physiol 63: 795-813. Atkinson L, Batten TF, Deuchars J. 2000. P2X(2) receptor immunoreactivity in the dorsal vagal complex and area postrema of the rat. Neuroscience 99: 683-696. Atkinson L, Batten TF, Moores TS, Varoqui H, Erickson JD, et al. 2004. Differential co‐localisation of the P2X7 receptor

subunit with vesicular glutamate transporters VGLUT1 and VGLUT2 in rat CNS. Neuroscience 123: 761-768. Bahima L, Aleu J, Elias M, Martin‐Satue M, Muhaisen A, et al. 2006. Endogenous hemichannels play a role in the release of ATP from Xenopus oocytes. J Cell Physiol 206: 95-102. Ballerini P, Di Iorio P, Ciccarelli R, Nargi E, D’Alimonte I, et al. 2002. Glial cells express multiple ATP binding cassette proteins which are involved in ATP release. Neuroreport 13: 1789-1792. Ballerini P, Rathbone MP, Di Iorio P, Renzetti A, Giuliani P, et al. 1996. Rat astroglial P2Z (P2X7) receptors regulate intracellular calcium and purine release. Neuroreport 7: 2533-2537. Barclay J, Patel S, Dorn G, Wotherspoon G, Moffatt S, et al. 2002. Functional downregulation of P2X3 receptor subunit in rat sensory neurons reveals a significant role in chronic neuropathic and inflammatory pain. J Neurosci 22: 8139-8147. Bardoni R, Goldstein PA, Lee CJ, Gu JG, MacDermott AB. 1997. ATP P2X receptors mediate fast synaptic transmission in the dorsal horn of the rat spinal cord. J Neurosci 17: 5297-5304. Barnard EA, Simon J. 2001. An elusive receptor is finally caught: P2Y(120 ), an important drug target in platelets. Trends Pharmacol Sci 22: 388-391. Bennett GC, Boarder MR. 2000. The effect of nucleotides and adenosine on stimulus‐evoked glutamate release from rat brain cortical slices. Br J Pharmacol 131: 617-623.

ATP‐mediated signaling in the nervous system Bianco F, Fumagalli M, Pravettoni E, D’Ambrosi N, Volonte C, et al. 2005. Pathophysiological roles of extracellular nucleotides in glial cells: Differential expression of purinergic receptors in resting and activated microglia. Brain Res Brain Res Rev 48: 144-156. Boarder MR, Webb TE. 2001. P2Y receptors: Structure and function. In: Handbook of experimental pharmacology: Purinergic and pyrimidiniergic signalling Vol. 151/I, Abbracchio MP, Williams M, editors. Berlin: Springer; pp. 65-89. Boehm S. 1999. ATP stimulates sympathetic transmitter release via presynaptic P2X purinoceptors. J Neurosci 19: 737-746. Boehm S, Kubista H. 2002. Fine tuning of sympathetic transmitter release via ionotropic and metabotropic presynaptic receptors. Pharmacol Rev 54: 43-99. Bonan CD, Walz R, Pereira GS, Worm PV, Battastini AM, et al. 2000. Changes in synaptosomal ectonucleotidase activities in two rat models of temporal lobe epilepsy. Epilepsy Res 39: 229-238. Bowser DN, Khakh BS. 2004. ATP excites interneurons and astrocytes to increase synaptic inhibition in neuronal networks. J Neurosci 24: 8606-8620. Bradbury EJ, Burnstock G, McMahon SB. 1998. The expression of P2X3 purinoreceptors in sensory neurons: Effects of axotomy and glial‐derived neurotrophic factor. Mol Cell Neurosci 12: 256-268. Braun N, Lenz C, Gillardon F, Zimmermann M, Zimmermann H. 1997. Focal cerebral ischemia enhances glial expression of ecto‐50 ‐nucleotidase. Brain Res 766: 213-226. Braun N, Zhu Y, Krieglstein J, Culmsee C, Zimmermann H. 1998. Upregulation of the enzyme chain hydrolyzing extracellular ATP after transient forebrain ischemia in the rat. J Neurosci 18: 4891-4900. Brough D, Le Feuvre RA, Iwakura Y, Rothwell NJ. 2002. Purinergic (P2X7) receptor activation of microglia induces cell death via an interleukin‐1‐independent mechanism. Mol Cell Neurosci 19: 272-280. Brown SG, Townsend‐Nicholson A, Jacobson KA, Burnstock G, King BF. 2002. Heteromultimeric P2X(1/2) receptors show a novel sensitivity to extracellular pH. J Pharmacol Exp Ther 300: 673-680. Burnstock G. 1972. Purinergic nerves. Pharmacol Rev 24: 509-581. Burnstock G. 2001. Purine‐mediated signalling in pain and visceral perception. Trends Pharmacol Sci 22: 182-188. Burnstock G. 2006. Purinergic P2 receptors as targets for novel analgesics. Pharmacol Ther 110(3): 433-454. Burnstock G, Wood JN. 1996. Purinergic receptors: Their role in nociception and primary afferent neurotransmission. Curr Opin Neurobiol 6: 526-532.

10

Calvert JA, Evans RJ. 2004. Heterogeneity of P2X receptors in sympathetic neurons: Contribution of neuronal P2X1 receptors revealed using knockout mice. Mol Pharmacol 65: 139-148. Cass CE, Young JD, Baldwin SA, Cabrita MA, Graham KA, et al. 1999. Nucleoside transporters of mammalian cells. Pharm Biotechnol 12: 313-352. Chakfe Y, Seguin R, Antel JP, Morissette C, Malo D, et al. 2002. ADP and AMP induce interleukin‐1beta release from microglial cells through activation of ATP‐primed P2X7 receptor channels. J Neurosci 22: 3061-3069. Chambers JK, Macdonald LE, Sarau HM, Ames RS, Freeman K, et al. 2000. A G protein‐coupled receptor for UDP‐ glucose. J Biol Chem 275: 10767-10771. Chessell IP, Michel AD, Humphrey PP. 1997. Properties of the pore‐forming P2X7 purinoceptor in mouse NTW8 microglial cells. Br J Pharmacol 121: 1429-1437. Chessell IP, Hatcher JP, Bountra C, Michel AD, Hughes JP, et al. 2005. Disruption of the P2X7 purinoceptor gene abolishes chronic inflammatory and neuropathic pain. Pain 114: 386-396. Chizh BA, Illes P. 2001. P2X receptors and nociception. Pharmacol Rev 53: 553-568. Choi HB, Hong SH, Ryu JK, Kim SU, McLarnon JG. 2003. Differential activation of subtype purinergic receptors modulates Ca(2þ) mobilization and COX‐2 in human microglia. Glia 43: 95-103. Cockayne DA, Dunn PM, Zhong Y, Rong W, Hamilton SG, et al. 2005. P2X2 knockout mice and P2X2/P2X3 double knockout mice reveal a role for the P2X2 receptor subunit in mediating multiple sensory effects of ATP. J Physiol 567: 621-639. Cockayne DA, Hamilton SG, Zhu QM, Dunn PM, Zhong Y, et al. 2000. Urinary bladder hyporeflexia and reduced pain‐ related behaviour in P2X3‐deficient mice. Nature 407: 1011-1015. Coco S, Calegari F, Pravettoni E, Pozzi D, Taverna E, et al. 2003. Storage and release of ATP from astrocytes in culture. J Biol Chem 278: 1354-1362. Collier HO, James GW, Schneider C. 1966. Antagonism by aspirin and fenamates of bronchoconstriction and nociception induced by adenosine‐50 ‐triphosphate. Nature 212: 411-412. Collo G, Neidhart S, Kawashima E, Kosco‐Vilbois M, North RA, et al. 1997. Tissue distribution of the P2X7 receptor. Neuropharmacology 36: 1277-1283. Collo G, North RA, Kawashima E, Merlo‐Pich E, Neidhart S, et al. 1996. Cloning of P2X5 and P2X6 receptors and the distribution and properties of an extended family of ATP‐gated ion channels. J Neurosci 16: 24952507.

247

248

10

ATP‐mediated signaling in the nervous system

Communi D, Gonzalez NS, Detheux M, Brezillon S, Lannoy V, et al. 2001. Identification of a novel human ADP receptor coupled to G(i). J Biol Chem 276: 41479-41485. Cook SP, McCleskey EW. 2002. Cell damage excites nociceptors through release of cytosolic ATP. Pain 95: 41-47. Cunha RA. 2001. Adenosine as a neuromodulator and as a homeostatic regulator in the nervous system: Different roles, different sources and different receptors. Neurochem Int 38: 107-125. Cunha RA, Ribeiro JA. 2000a. ATP as a presynaptic modulator. Life Sci 68: 119-137. Cunha RA, Ribeiro JA. 2000b. Purinergic modulation of [(3)H]GABA release from rat hippocampal nerve terminals. Neuropharmacology 39: 1156-1167. Cunha RA, Ribeiro JA, Sebastiao AM. 1994. Purinergic modulation of the evoked release of [3H]acetylcholine from the hippocampus and cerebral cortex of the rat: Role of the ectonucleotidases. Eur J Neurosci 6: 33-42. Cunha RA, Sebastiao AM, Ribeiro JA. 1992. Ecto‐50 ‐nucleotidase is associated with cholinergic nerve terminals in the hippocampus but not in the cerebral cortex of the rat. J Neurochem 59: 657-666. Cunha RA, Sebastiao AM, Ribeiro JA. 1998. Inhibition by ATP of hippocampal synaptic transmission requires localized extracellular catabolism by ecto‐nucleotidases into adenosine and channeling to adenosine A1 receptors. J Neurosci 18: 1987-1995. Darby M, Kuzmiski JB, Panenka W, Feighan D, MacVicar BA. 2003. ATP released from astrocytes during swelling activates chloride channels. J Neurophysiol 89: 1870-1877. Deuchars SA, Atkinson L, Brooke RE, Musa H, Milligan CJ, et al. 2001. Neuronal P2X7 receptors are targeted to presynaptic terminals in the central and peripheral nervous systems. J Neurosci 21: 7143-7152. Dorn G, Patel S, Wotherspoon G, Hemmings‐Mieszczak M, Barclay J, et al. 2004. siRNA relieves chronic neuropathic pain. Nucleic Acids Res 32: e49. Drury AN, Szent‐Gyo¨rgyi A. 1929. The physiological action of adenine compounds with especial reference to their action on the mammalian heart. J Physiol (Lond) 68: 214-237. Dulla CG, Dobelis P, Pearson T, Frenguelli BG, Staley KJ, et al. 2005. Adenosine and ATP link PCO2 to cortical excitability via pH. Neuron 48: 1011-1023. Dunn PM, Zhong Y, Burnstock G. 2001. P2X receptors in peripheral neurons. Prog Neurobiol 65: 107-134. Dunwiddie TV, Diao L, Proctor WR. 1997. Adenine nucleotides undergo rapid, quantitative conversion to adenosine in the extracellular space in rat hippocampus. J Neurosci 17: 7673-7682. Edwards FA, Gibb AJ, Colquhoun D. 1992. ATP receptor‐ mediated synaptic currents in the central nervous system. Nature 359: 144-147.

Egan TM, Khakh BS. 2004. Contribution of calcium ions to P2X channel responses. J Neurosci 24: 3413-3420. Evans RJ, Derkach V, Surprenant A. 1992. ATP mediates fast synaptic transmission in mammalian neurons. Nature 357: 503-505. Fellin T, Pozzan T, Carmignoto G. 2006. Purinergic receptors mediate two distinct glutamate release pathways in hippocampal astrocytes. J Biol Chem. 281: 4274-4284. Ferrari D, Chiozzi P, Falzoni S, Hanau S, Di Virgilio F. 1997c. Purinergic modulation of interleukin‐1 beta release from microglial cells stimulated with bacterial endotoxin. J Exp Med 185: 579-582. Ferrari D, Chiozzi P, Falzoni S, Dal Susino M, Collo G, et al. 1997a. ATP‐mediated cytotoxicity in microglial cells. Neuropharmacology 36: 1295-1301. Ferrari D, Chiozzi P, Falzoni S, Dal Susino M, Melchiorri L, et al. 1997b. Extracellular ATP triggers IL‐1 beta release by activating the purinergic P2Z receptor of human macrophages. J Immunol 159: 1451-1458. Ferrari D, Los M, Bauer MK, Vandenabeele P, Wesselborg S, et al. 1999. P2Z purinoreceptor ligation induces activation of caspases with distinct roles in apoptotic and necrotic alterations of cell death. FEBS Lett 447: 71-75. Ferrari D, Villalba M, Chiozzi P, Falzoni S, Ricciardi‐Castagnoli P, et al. 1996. Mouse microglial cells express a plasma membrane pore gated by extracellular ATP. J Immunol 156: 1531-1539. Fields RD, Stevens B. 2000. ATP: An extracellular signaling molecule between neurons and glia. Trends Neurosci 23: 625-633. Franke H, Illes P. 2006. Involvement of P2 receptors in the growth and survival of neurons in the CNS. Pharmacol Ther 109(3): 297-324. Galligan JJ, Bertrand PP. 1994. ATP mediates fast synaptic potentials in enteric neurons. J Neurosci 14: 7563-7571. Gendron FP, Chalimoniuk M, Strosznajder J, Shen S, Gonzalez FA, et al. 2003. P2X7 nucleotide receptor activation enhances IFN gamma‐induced type II nitric oxide synthase activity in BV‐2 microglial cells. J Neurochem 87: 344-352. Gerevich Z, Borvendeg SJ, Schroder W, Franke H, Wirkner K, et al. 2004. Inhibition of N‐type voltage‐activated calcium channels in rat dorsal root ganglion neurons by P2Y receptors is a possible mechanism of ADP‐induced analgesia. J Neurosci 24: 797-807. Gomes P, Srinivas SP, Van Driessche W, Vereecke J, Himpens B. 2005. ATP release through connexin hemichannels in corneal endothelial cells. Invest Ophthalmol Vis Sci 46: 1208-1218. Gordon GR, Baimoukhametova DV, Hewitt SA, Rajapaksha WR, Fisher TE, et al. 2005. Norepinephrine triggers release of glial ATP to increase postsynaptic efficacy. Nat Neurosci 8: 1078-1086.

ATP‐mediated signaling in the nervous system Gourine AV. 2005. On the peripheral and central chemoreception and control of breathing: An emerging role of ATP. J Physiol 568: 715-724. Gourine AV, Llaudet E, Dale N, Spyer KM. 2005a. ATP is a mediator of chemosensory transduction in the central nervous system. Nature 436: 108-111. Gourine AV, Llaudet E, Dale N, Spyer KM. 2005b. Release of ATP in the ventral medulla during hypoxia in rats: Role in hypoxic ventilatory response. J Neurosci 25: 1211-1218. Griffiths M, Yao SY, Abidi F, Phillips SE, Cass CE, et al. 1997. Molecular cloning and characterization of a nitrobenzylthioinosine‐insensitive (ei) equilibrative nucleoside transporter from human placenta. Biochem J 328(Pt 3): 739-743. Grondal EJ, Janetzko A, Zimmermann H. 1988. Monospecific antiserum against 50 ‐nucleotidase from Torpedo electric organ: Immunocytochemical distribution of the enzyme and its association with Schwann cell membranes. Neuroscience 24: 351-363. Gu JG, MacDermott AB. 1997. Activation of ATP P2X receptors elicits glutamate release from sensory neuron synapses. Nature 389: 749-753. Guo LH, Trautmann K, Schluesener HJ. 2005. Expression of P2X4 receptor by lesional activated microglia during formalin‐induced inflammatory pain. J Neuroimmunol 163: 120-127. Guthrie PB, Knappenberger J, Segal M, Bennett MV, Charles AC, et al. 1999. ATP released from astrocytes mediates glial calcium waves. J Neurosci 19: 520-528. Haas S, Brockhaus J, Verkhratsky A, Kettenmann H. 1996. ATP‐induced membrane currents in ameboid microglia acutely isolated from mouse brain slices. Neuroscience 75: 257-261. Hamann M, Attwell D. 1996. Non‐synaptic release of ATP by electrical stimulation in slices of rat hippocampus, cerebellum and habenula. Eur J Neurosci 8: 1510-1515. Harden TK, Lazarowski ER, Boucher RC. 1997. Release, metabolism and interconversion of adenine and uridine nucleotides: Implications for G protein‐coupled P2 receptor agonist selectivity. Trends Pharmacol Sci 18: 43-46. Hide I, Tanaka M, Inoue A, Nakajima K, Kohsaka S, et al. 2000. Extracellular ATP triggers tumor necrosis factor‐ alpha release from rat microglia. J Neurochem 75: 965-972. Hollopeter G, Jantzen HM, Vincent D, Li G, England L, et al. 2001. Identification of the platelet ADP receptor targeted by antithrombotic drugs. Nature 409: 202-207. Holton P. 1959. The liberation of adenosine triphosphate on antidromic stimulation of sensory nerves. J Physiol 145: 494-504. Honore P, Kage K, Mikusa J, Watt AT, Johnston JF, et al. 2002. Analgesic profile of intrathecal P2X(3) antisense oligonucleotide treatment in chronic inflammatory and neuropathic pain states in rats. Pain 99: 11-19.

10

Hugel S, Schlichter R. 2000. Presynaptic P2X receptors facilitate inhibitory GABAergic transmission between cultured rat spinal cord dorsal horn neurons. J Neurosci 20: 2121-2130. Illes P, Alexandre Ribeiro J. 2004. Molecular physiology of P2 receptors in the central nervous system. Eur J Pharmacol 483: 5-17. Illes P, Klotz K‐N, Lohse MJ. 2000. Signaling by extracellular nucleotides and nucleosides. Naunyn Schmiedeberg’s Arch Pharmacol 362: 295-299. Inoue K. 2002. Microglial activation by purines and pyrimidines. Glia 40: 156-163. Inoue K, Tsuda M, Koizumi S. 2004. ATP‐ and adenosine‐ mediated signaling in the central nervous system: Chronic pain and microglia: Involvement of the ATP receptor P2X4. J Pharmacol Sci 94: 112-114. Inoue K, Koizumi S, Ueno S, Kita A, Tsuda M. 1999. The functions of ATP receptors in the synaptic transmission in the hippocampus. Prog Brain Res 120: 193-206. Inoue K, Nakajima K, Morimoto T, Kikuchi Y, Koizumi S, et al. 1998. ATP stimulation of Ca2þ‐dependent plasminogen release from cultured microglia. Br J Pharmacol 123: 1304-1310. James S, Richardson PJ. 1993. Production of adenosine from extracellular ATP at the striatal cholinergic synapse. J Neurochem 60: 219-227. Jo YH, Role LW. 2002. Coordinate release of ATP and GABA at in vitro synapses of lateral hypothalamic neurons. J Neurosci 22: 4794-4804. Jones CA, Vial C, Sellers LA, Humphrey PP, Evans RJ, et al. 2004. Functional regulation of P2X6 receptors by N‐linked glycosylation: Identification of a novel alpha beta‐methylene ATP‐sensitive phenotype. Mol Pharmacol 65: 979-985. Juranyi Z, Orso E, Janossy A, Szalay KS, Sperlagh B, et al. 1997. ATP and [3H]noradrenaline release and the presence of ecto‐Ca(2þ)‐ATPases in the capsule‐glomerulosa fraction of the rat adrenal gland. J Endocrinol 153: 105114. Kanjhan R, Housley GD, Burton LD, Christie DL, Kippenberger A, et al. 1999. Distribution of the P2X2 receptor subunit of the ATP‐gated ion channels in the rat central nervous system. J Comp Neurol 407: 11-32. Kato F, Shigetomi E. 2001. Distinct modulation of evoked and spontaneous EPSCs by purinoceptors in the nucleus tractus solitarii of the rat. J Physiol 530: 469-486. Kegel B, Braun N, Heine P, Maliszewski CR, Zimmermann H. 1997. An ecto‐ATPase and an ecto‐ATP diphosphohydrolase are expressed in rat brain. Neuropharmacology 36: 1189-1200. Kennedy C. 2001. The role of purines in the peripheral nervous system. In: Purinergic and pyrimidinergic signalling I. Molecular, Nervous and Urogenitary sytem function,

249

250

10

ATP‐mediated signaling in the nervous system

Vol. 151. Abbracchio MP, Williams M, editors. Berlin: Springer; pp. 289-305. Kennedy C. 2005. P2X receptors: Targets for novel analgesics? Neuroscientist 11: 345-356. Kennedy C, Assis TS, Currie AJ, Rowan EG. 2003. Crossing the pain barrier: P2 receptors as targets for novel analgesics. J Physiol 553: 683-694. Khakh BS. 2001. Molecular physiology of P2X receptors and ATP signalling at synapses. Nat Rev Neurosci 2: 165-174. Khakh BS, Henderson G. 1998. ATP receptor‐mediated enhancement of fast excitatory neurotransmitter release in the brain. Mol Pharmacol 54: 372-378. Khakh BS, Gittermann D, Cockayne DA, Jones A. 2003. ATP modulation of excitatory synapses onto interneurons. J Neurosci 23: 7426-7437. Khakh BS, Burnstock G, Kennedy C, King BF, North A, Se´guela P, et al. 2001. International union of pharmacology. XXIV. Current status of the nomenclature and properties of P2x receptors and their subunits. Pharmacol Rev 53: 107118. King BF, Townsend‐Nicholson A, Wildman SS, Thomas T, Spyer KM, et al. 2000. Coexpression of rat P2X2 and P2X6 subunits in Xenopus oocytes. J Neurosci 20: 4871-4877. Kirkpatrick K, Burnstock G. 1987. Sympathetic nerve‐ mediated release of ATP from the guinea‐pig vas deferens is unaffected by reserpine. Eur J Pharmacol 138: 207-214. Kittner H, Krugel U, Illes P. 2001. The purinergic P2 receptor antagonist pyridoxalphosphate‐6‐azophenyl‐20 40 ‐ disulphonic acid prevents both the acute locomotor effects of amphetamine and the behavioural sensitization caused by repeated amphetamine injections in rats. Neuroscience 102: 241-243. Kittner H, Krugel U, Hoffmann E, Illes P. 2000. Effects of intra‐accumbens injection of 2‐methylthio ATP: A combined open field and electroencephalographic study in rats. Psychopharmacology (Berl) 150: 123-131. Kittner H, Krugel U, Hoffmann E, Illes P. 2004. Modulation of feeding behaviour by blocking purinergic receptors in the rat nucleus accumbens: A combined microdialysis, electroencephalographic and behavioural study. Eur J Neurosci 19: 396-404. Kittner H, Franke H, Fischer W, Schultheis N, Krugel U, et al. 2003. Stimulation of P2Y1 receptors causes anxiolytic‐like effects in the rat elevated plus‐maze: Implications for the involvement of P2Y1 receptor‐mediated nitric oxide production. Neuropsychopharmacology 28: 435-444. Knott TK, Velazquez‐Marrero C, Lemos JR. 2005. ATP elicits inward currents in isolated vasopressinergic neurohypophysial terminals via P2X2 and P2X3 receptors. Pflugers Arch 450: 381-389.

Koch H, Kugelgen I, Starke K. 1997. P2‐receptor‐mediated inhibition of noradrenaline release in the rat hippocampus. Naunyn Schmiedebergs Arch Pharmacol 355: 707-715. Koizumi S, Inoue K. 1997. Inhibition by ATP of calcium oscillations in rat cultured hippocampal neurones. Br J Pharmacol 122: 51-58. Koizumi S, Fujishita K, Tsuda M, Shigemoto‐Mogami Y, Inoue K. 2003. Dynamic inhibition of excitatory synaptic transmission by astrocyte‐derived ATP in hippocampal cultures. Proc Natl Acad Sci USA 100: 11023-11028. Koles L, Furst S, Illes P. 2005. P2X and P2Y receptors as possible targets of therapeutic manipulations in CNS illnesses. Drug News Perspect 18: 85-101. Krugel U, Kittner H, Illes P. 1999. Adenosine 50 ‐triphosphate‐ induced dopamine release in the rat nucleus accumbens in vivo. Neurosci Lett 265: 49-52. Krugel U, Kittner H, Illes P. 2001. Mechanisms of adenosine 50 ‐triphosphate‐induced dopamine release in the rat nucleus accumbens in vivo. Synapse 39: 222-232. Kukley M, Stausberg P, Adelmann G, Chessell IP, Dietrich D. 2004. Ecto‐nucleotidases and nucleoside transporters mediate activation of adenosine receptors on hippocampal mossy fibers by P2X7 receptor agonist 20 ‐30 ‐O‐(4‐benzoylbenzoyl)‐ATP. J Neurosci 24: 7128-7139. Latini S, Pedata F. 2001. Adenosine in the central nervous system: Release mechanisms and extracellular concentrations. J Neurochem 79: 463-484. Le KT, Babinski K, Seguela P. 1998. Central P2X4 and P2X6 channel subunits coassemble into a novel heteromeric ATP receptor. J Neurosci 18: 7152-7159. Le KT, Paquet M, Nouel D, Babinski K, Seguela P. 1997. Primary structure and expression of a naturally truncated human P2X ATP receptor subunit from brain and immune system. FEBS Lett 418: 195-199. Lee CW, Jarvis SM. 1988. Nucleoside transport in rat cerebral‐ cortical synaptosomes. Evidence for two types of nucleoside transporters. Biochem J 249: 557-564. Lewis C, Neidhart S, Holy C, North RA, Buell G, et al. 1995. Coexpression of P2X2 and P2X3 receptor subunits can account for ATP‐gated currents in sensory neurons [see comments]. Nature 377: 432-435. Li C, Peoples RW, Lanthorn TH, Li ZW, Weight FF. 1999. Distinct ATP‐activated currents in different types of neurons dissociated from rat dorsal root ganglion. Neurosci Lett 263: 57-60. Llaudet E, Hatz S, Droniou M, Dale N. 2005. Microelectrode biosensor for real‐time measurement of ATP in biological tissue. Anal Chem 77: 3267-3273. MacDermott AB, Role LW, Siegelbaum SA. 1999. Presynaptic ionotropic receptors and the control of transmitter release. Annu Rev Neurosci 22: 443-485.

ATP‐mediated signaling in the nervous system McGehee DS, Role LW. 1996. Presynaptic ionotropic receptors. Curr Opin Neurobiol 6: 342-349. McLarnon JG. 2005. Purinergic mediated changes in Ca2þ mobilization and functional responses in microglia: Effects of low levels of ATP. J Neurosci Res 81: 349-356. Mendoza‐Fernandez V, Andrew RD, Barajas‐Lopez C. 2000. ATP inhibits glutamate synaptic release by acting at P2Y receptors in pyramidal neurons of hippocampal slices. J Pharmacol Exp Ther 293: 172-179. Moore DJ, Chambers JK, Wahlin JP, Tan KB, Moore GB, et al. 2001. Expression pattern of human P2Y receptor subtypes: A quantitative reverse transcription‐polymerase chain reaction study. Biochim Biophys Acta 1521: 107-119. Moran‐Jimenez MJ, Matute C. 2000. Immunohistochemical localization of the P2Y(1) purinergic receptor in neurons and glial cells of the central nervous system. Brain Res Mol Brain Res 78: 50-58. Nagy AK, Shuster TA, Delgado‐Escueta AV. 1989. Rat brain synaptosomal ATP:AMP‐phosphotransferase activity. J Neurochem 53: 1166-1172. Nakatsuka T, Gu JG. 2001. ATP P2X receptor‐mediated enhancement of glutamate release and evoked EPSCs in dorsal horn neurons of the rat spinal cord. J Neurosci 21: 6522-6531. Nakatsuka T, Tsuzuki K, Ling JX, Sonobe H, Gu JG. 2003. Distinct roles of P2X receptors in modulating glutamate release at different primary sensory synapses in rat spinal cord. J Neurophysiol 89: 3243-3252. Newman EA. 2003a. Glial cell inhibition of neurons by release of ATP. J Neurosci 23: 1659-1666. Newman EA. 2003b. New roles for astrocytes: Regulation of synaptic transmission. Trends Neurosci 26: 536-542. Nicholas RA. 2001. Identification of the P2Y(12) receptor: A novel member of the P2Y family of receptors activated by extracellular nucleotides. Mol Pharmacol 60: 416-420. Nicke A, Kerschensteiner D, Soto F. 2005. Biochemical and functional evidence for heteromeric assembly of P2X1 and P2X4 subunits. J Neurochem 92: 925-933. Nieber K, Poelchen W, Illes P. 1997. Role of ATP in fast excitatory synaptic potentials in locus coeruleus neurones of the rat. Br J Pharmacol 122: 423-430. Norenberg W, Langosch JM, Gebicke‐Haerter PJ, Illes P. 1994. Characterization and possible function of adenosine 50 ‐triphosphate receptors in activated rat microglia. Br J Pharmacol 111: 942-950. North RA, Surprenant A. 2000. Pharmacology of cloned P2X receptors. Annu Rev Pharmacol Toxicol 40: 563-580. Ogata T, Chuai M, Morino T, Yamamoto H, Nakamura Y, et al. 2003. Adenosine triphosphate inhibits cytokine release from lipopolysaccharide‐activated microglia via P2y receptors. Brain Res 981: 174-183.

10

Ohtani Y, Minami M, Satoh M. 2000. Expression of inducible nitric oxide synthase mRNA and production of nitric oxide are induced by adenosine triphosphate in cultured rat microglia. Neurosci Lett 293: 72-74. Okada M, Kawata Y, Murakami T, Wada K, Mizuno K, et al. 1999. Interaction between purinoceptor subtypes on hippocampal serotonergic transmission using in vivo microdialysis. Neuropharmacology 38: 707-715. Pankratov Y, Castro E, Miras‐Portugal MT, Krishtal O. 1998. A purinergic component of the excitatory postsynaptic current mediated by P2X receptors in the CA1 neurons of the rat hippocampus. Eur J Neurosci 10: 3898-3902. Pankratov Y, Lalo U, Krishtal O, Verkhratsky A. 2002. Ionotropic P2X purinoreceptors mediate synaptic transmission in rat pyramidal neurones of layer II/III of somato‐sensory cortex. J Physiol 542: 529-536. Pascual O, Casper KB, Kubera C, Zhang J, Revilla‐Sanchez R, et al. 2005. Astrocytic purinergic signaling coordinates synaptic networks. Science 310: 113-116. Papp L, Vizi ES, Sperlagh B. 2004b. Lack of ATP‐evoked GABA and glutamate release in the hippocampus of P2X7 receptor/ mice. Neuroreport 15: 2387-2391. Papp L, Balazsa T, Kofalvi A, Erdelyi F, Szabo G, et al. 2004a. P2X receptor activation elicits transporter‐mediated noradrenaline release from rat hippocampal slices. J Pharmacol Exp Ther 310: 973-980. Parvathenani LK, Tertyshnikova S, Greco CR, Roberts SB, Robertson B, et al. (2003) P2X7 mediates superoxide production in primary microglia and is up‐regulated in a transgenic mouse model of Alzheimer’s disease. J Biol Chem 278: 13309-13317. Patel MK, Khakh BS, Henderson G. 2001. Properties of native P2X receptors in rat trigeminal mesencephalic nucleus neurones: Lack of correlation with known, heterologously expressed P2X receptors. Neuropharmacology 40: 96-105. Pearson RA, Dale N, Llaudet E, Mobbs P. 2005. ATP released via gap junction hemichannels from the pigment epithelium regulates neural retinal progenitor proliferation. Neuron 46: 731-744. Petruska JC, Cooper BY, Gu JG, Rau KK, Johnson RD. 2000. Distribution of P2X1, P2X2, and P2X3 receptor subunits in rat primary afferents: Relation to population markers and specific cell types. J Chem Neuroanat 20: 141-162. Poelchen W, Sieler D, Wirkner K, Illes P. 2001. Co‐transmitter function of ATP in central catecholaminergic neurons of the rat. Neuroscience 102: 593-602. Queiroz G, Talaia C, Goncalves J. 2003. ATP modulates noradrenaline release by activation of inhibitory P2Y receptors and facilitatory P2X receptors in the rat vas deferens. J Pharmacol Exp Ther 307: 809-815.

251

252

10

ATP‐mediated signaling in the nervous system

Ralevic V, Burnstock G. 1998. Receptors for purines and pyrimidines. Pharmacol Rev 50: 413-492. Rampe D, Wang L, Ringheim GE. 2004. P2X7 receptor modulation of beta‐amyloid‐ and LPS‐induced cytokine secretion from human macrophages and microglia. J Neuroimmunol 147: 56-61. Rhee JS, Wang ZM, Nabekura J, Inoue K, Akaike N. 2000. ATP facilitates spontaneous glycinergic IPSC frequency at dissociated rat dorsal horn interneuron synapses. J Physiol 524 Pt 2: 471-483. Rodrigues RJ, Almeida T, Richardson PJ, Oliveira CR, Cunha RA. 2005. Dual presynaptic control by ATP of glutamate release via facilitatory P2X1, P2X2/3, and P2X3 and inhibitory P2Y1, P2Y2, and/or P2Y4 receptors in the rat hippocampus. J Neurosci 25: 6286-6295. Rogers M, Colquhoun LM, Patrick JW, Dani JA. 1997. Calcium flux through predominantly independent purinergic ATP and nicotinic acetylcholine receptors. J Neurophysiol 77: 1407-1417. Rong W, Gourine AV, Cockayne DA, Xiang Z, Ford AP, et al. 2003. Pivotal role of nucleotide P2X2 receptor subunit of the ATP‐gated ion channel mediating ventilatory responses to hypoxia. J Neurosci 23: 11315-11321. Ruan HZ, Burnstock G. 2003. Localisation of P2Y1 and P2Y4 receptors in dorsal root, nodose and trigeminal ganglia of the rat. Histochem Cell Biol 120: 415-426. Ruan HZ, Birder LA, de Groat WC, Tai C, Roppolo J, et al. 2005. Localization of P2X and P2Y receptors in dorsal root ganglia of the cat. J Histochem Cytochem 53: 1273-1282. Rubio M, Soto F. 2001. Distinct Localization of P2X receptors at excitatory postsynaptic specializations. J Neurosci 21: 641-653. Sanchez‐Nogueiro J, Marin‐Garcia P, Miras‐Portugal MT. 2005. Characterization of a functional P2X(7)‐like receptor in cerebellar granule neurons from P2X(7) knockout mice. FEBS Lett 579: 3783-3788. Sanz JM, Di Virgilio F. 2000. Kinetics and mechanism of ATP‐ dependent IL‐1 beta release from microglial cells. J Immunol 164: 4893-4898. Schoen SW, Ebert U, Loscher W. 1999. 50 ‐Nucleotidase activity of mossy fibers in the dentate gyrus of normal and epileptic rats. Neuroscience 93: 519-526. SchoenSW, Graeber MB, Reddington M, Kreutzberg GW. 1987. Light and electron microscopical immunocytochemistry of 50 ‐nucleotidase in rat cerebellum. Histochemistry 87: 107-113. Schwab JM, Guo L, Schluesener HJ. 2005. Spinal cord injury induces early and persistent lesional P2X4 receptor expression. J Neuroimmunol 163: 185-189. Schwiebert EM. 1999. ABC transporter‐facilitated ATP conductive transport. Am J Physiol 276: C1-8.

Shigetomi E, Kato F. 2004. Action potential‐independent release of glutamate by Ca2þ entry through presynaptic P2X receptors elicits postsynaptic firing in the brainstem autonomic network. J Neurosci 24: 3125-3135. Shinozuka K, Mizuno H, Nakamura K, Kunitomo M. 2002. Purinergic modulation of vascular sympathetic neurotransmission. Jpn J Pharmacol 88: 19-25. Silinsky EM, Gerzanich V, Vanner SM. 1992. ATP mediates excitatory synaptic transmission in mammalian neurones. Br J Pharmacol 106: 762-763. Sim JA, Young MT, Sung HY, North RA, Surprenant A. 2004. Reanalysis of P2X7 receptor expression in rodent brain. J Neurosci 24: 6307-6314. Smith DO. 1991. Sources of adenosine released during neuromuscular transmission in the rat. J Physiol 432: 343-354. Sneddon P, Westfall TD, Todorov LD, Todorova SM, Westfall DP, et al. 2000. The effect of P2 receptor antagonists and ATPase inhibition on sympathetic purinergic neurotransmission in the guinea‐pig isolated vas deferens. Br J Pharmacol 129: 1089-1094. Souslova V, Cesare P, Ding Y, Akopian AN, Stanfa L, et al. 2000. Warm‐coding deficits and aberrant inflammatory pain in mice lacking P2X3 receptors. Nature 407: 1015-1017. Sperla´gh B. Vizi ES. 1996. Neuronal synthesis, storage and release of ATP. Seminars in the Neurosciences 8: 175-186. Sperlagh B, Vizi ES. 1991. Effect of presynaptic P2 receptor stimulation on transmitter release. J Neurochem 56: 1466-1470. Sperlagh B, Vizi ES. 1992. Is the neuronal ATP release from guinea‐pig vas deferens subject to alpha 2‐adrenoceptor‐ mediated modulation? Neuroscience 51: 203-209. Sperlagh B, Vizi ES. 2000. Regulation of purine release. In: Handbook of experimental pharmacology, Vol. 151. Abbracchio MP, Williams M, editors. Springer; pp. 179-209. Sperlagh B, Andras I, Vizi S. 1997. Effect of subtype‐ specific Ca(2þ)‐antagonists and Ca(2þ)‐free media on the field stimulation‐evoked release of ATP and [3H]acetylcholine from rat habenula slices. Neurochem Res 22: 967-975. Sperlagh B, Erdelyi F, Szabo G, Vizi ES. 2000. Local regulation of [(3)H]‐noradrenaline release from the isolated guinea‐ pig right atrium by P(2X)‐receptors located on axon terminals. Br J Pharmacol 131: 1775-1783. Sperlagh B, Kittel A, Lajtha A, Vizi ES. 1995. ATP acts as fast neurotransmitter in rat habenula: Neurochemical and enzymecytochemical evidence. Neuroscience 66: 915-920. Sperlagh B, Magloczky Z, Vizi ES, Freund TF. 1998a. The triangular septal nucleus as the major source of ATP release in the rat habenula: A combined neurochemical and morphological study. Neuroscience 86: 1195-1207.

ATP‐mediated signaling in the nervous system Sperlagh B, Sershen H, Lajtha A, Vizi ES. 1998b. Co‐release of endogenous ATP and [3H]noradrenaline from rat hypothalamic slices: Origin and modulation by alpha2‐adrenoceptors. Neuroscience 82: 511-520. Sperlagh B, Mergl Z, Juranyi Z, Vizi ES, Makara GB. 1999. Local regulation of vasopressin and oxytocin secretion by extracellular ATP in the isolated posterior lobe of the rat hypophysis. J Endocrinol 160: 343-350. Sperlagh B, Szabo G, Erdelyi F, Baranyi M, Vizi ES. 2003. Homo‐ and heteroexchange of adenine nucleotides and nucleosides in rat hippocampal slices by the nucleoside transport system. Br J Pharmacol 139: 623-633. Sperlagh B, Kofalvi A, Deuchars J, Atkinson L, Milligan CJ, et al. 2002. Involvement of P2X7 receptors in the regulation of neurotransmitter release in the rat hippocampus. J Neurochem 81: 1196-1211. Spyer KM, Dale N, Gourine AV. 2004. ATP is a key mediator of central and peripheral chemosensory transduction. Exp Physiol 89: 53-59. Stout C, Goodenough DA, Paul DL. 2004. Connexins: Functions without junctions. Curr Opin Cell Biol 16: 507-512. Stout CE, Costantin JL, Naus CC, Charles AC. 2002. Intercellular calcium signaling in astrocytes via ATP release through connexin hemichannels. J Biol Chem 277: 10482-10488. Sun XP, Stanley EF. 1996. An ATP‐activated, ligand‐gated ion channel on a cholinergic presynaptic nerve terminal. Proc Natl Acad Sci USA 93: 1859-1863. Suzuki T, Hide I, Ido K, Kohsaka S, Inoue K, et al. 2004. Production and release of neuroprotective tumor necrosis factor by P2X7 receptor‐activated microglia. J Neurosci 24: 1-7. Terrian DM, Hernandez PG, Rea MA, Peters RI. 1989. ATP release, adenosine formation, and modulation of dynorphin and glutamic acid release by adenosine analogues in rat hippocampal mossy fiber synaptosomes. J Neurochem 53: 1390-1399. Thomas T, Spyer KM. 2000. ATP as a mediator of mammalian central CO2 chemoreception. J Physiol 523 Pt 2: 441-447. Thomas T, Ralevic V, Gadd CA, Spyer KM. 1999. Central CO2 chemoreception: A mechanism involving P2 purinoceptors localized in the ventrolateral medulla of the anaesthetized rat. J Physiol 517(Pt 3): 899-905. Thomas T, Ralevic V, Bardini M, Burnstock G, Spyer KM. 2001. Evidence for the involvement of purinergic signalling in the control of respiration. Neuroscience 107: 481-490. Thorn JA, Jarvis SM. 1996. Adenosine transporters. Gen Pharmacol 27: 613-620. Torres GE, Egan TM, Voigt MM. 1999. Hetero‐oligomeric assembly of P2X receptor subunits. Specificities exist with regard to possible partners. J Biol Chem 274: 6653-6659.

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Torres GE, Haines WR, Egan TM, Voigt MM. 1998. Co‐ expression of P2X1 and P2X5 receptor subunits reveals a novel ATP‐gated ion channel. Mol Pharmacol 54: 989-993. Trendelenburg AU, Bultmann R. 2000. P2 receptor‐mediated inhibition of dopamine release in rat neostriatum. Neuroscience 96: 249-252. Tsuda M, Shigemoto‐Mogami Y, Koizumi S, Mizokoshi A, Kohsaka S, et al. 2003. P2X4 receptors induced in spinal microglia gate tactile allodynia after nerve injury. Nature 424: 778-783. Tsuzuki K, Ase A, Seguela P, Nakatsuka T, Wang CY, et al. 2003. TNP‐ATP‐resistant P2X ionic current on the central terminals and somata of rat primary sensory neurons. J Neurophysiol 89: 3235-3242. Valera S, Hussy N, Evans RJ, Adami N, North RA, et al. 1994. A new class of ligand‐gated ion channel defined by P2x receptor for extracellular ATP. Nature 371: 516-519. Verderio C, Matteoli M. 2001. ATP mediates calcium signaling between astrocytes and microglial cells: Modulation by IFN‐gamma. J Immunol 166: 6383-6391. Visentin S, Renzi M, Frank C, Greco A, Levi G. 1999. Two different ionotropic receptors are activated by ATP in rat microglia. J Physiol 519 Pt 3: 723-736. Vizi ES. 2000. Role of high‐affinity receptors and membrane transporters in nonsynaptic communication and drug action in the central nervous system. Pharmacol Rev 52: 63-89. Vizi ES, Sperlagh B. 1999. Receptor‐ and carrier‐mediated release of ATP of postsynaptic origin: Cascade transmission. Prog Brain Res 120: 159-169. Vizi ES, Sperlagh B, Baranyi M. 1992. Evidence that ATP released from the postsynaptic site by noradrenaline, is involved in mechanical responses of guinea‐pig vas deferens: Cascade transmission. Neuroscience 50: 455-465. Vizi ES, Nitahara K, Sato K, Sperlagh B. 2000. Stimulation‐ dependent release, breakdown, and action of endogenous ATP in mouse hemidiaphragm preparation: The possible role of ATP in neuromuscular transmission. J Auton Nerv Syst 81: 278-284. Vizi ES, Liang SD, Sperlagh B, Kittel A, Juranyi Z. 1997. Studies on the release and extracellular metabolism of endogenous ATP in rat superior cervical ganglion: Support for neurotransmitter role of ATP. Neuroscience 79: 893-903. Vlaskovska M, Kasakov L, Rong W, Bodin P, Bardini M, et al. 2001. P2X3 knock‐out mice reveal a major sensory role for urothelially released ATP. J Neurosci 21: 5670-5677. von Kugelgen I, Wetter A. 2000. Molecular pharmacology of P2Y‐receptors. Naunyn Schmiedebergs Arch Pharmacol 362: 310-323. von Kugelgen I, Koch H, Starke K. 1997. P2‐receptor‐ mediated inhibition of serotonin release in the rat brain cortex. Neuropharmacology 36: 1221-1227.

253

254

10

ATP‐mediated signaling in the nervous system

von Kugelgen I, Spath L, Starke K. 1994. Evidence for P2‐ purinoceptor‐mediated inhibition of noradrenaline release in rat brain cortex. Br J Pharmacol 113: 815-822. Vulchanova L, Arvidsson U, Riedl M, Wang J, Buell G, et al. 1996. Differential distribution of two ATP‐gated channels (P2X receptors) determined by immunocytochemistry. Proc Natl Acad Sci USA 93: 8063-8067. Vulchanova L, Riedl MS, Shuster SJ, Buell G, Surprenant A, et al. 1997. Immunohistochemical study of the P2X2 and P2X3 receptor subunits in rat and monkey sensory neurons and their central terminals. Neuropharmacology 36: 1229-1242. Walter L, Dinh T, Stella N. 2004. ATP induces a rapid and pronounced increase in 2‐arachidonoylglycerol production by astrocytes, a response limited by monoacylglycerol lipase. J Neurosci 24: 8068-8074. Walz W, Ilschner S, Ohlemeyer C, Banati R, Kettenmann H. 1993. Extracellular ATP activates a cation conductance and a Kþ conductance in cultured microglial cells from mouse brain. J Neurosci 13: 4403-4411. Wang TF, Guidotti G. 1998. Widespread expression of ecto‐ apyrase (CD39) in the central nervous system. Brain Res 790: 318-322. Wang TF, Rosenberg PA, Guidotti G. 1997. Characterization of brain ecto‐apyrase: Evidence for only one ecto‐apyrase (CD39) gene. Brain Res Mol Brain Res 47: 295-302. Wang Y, Roman R, Lidofsky SD, Fitz JG. 1996. Autocrine signaling through ATP release represents a novel mechanism for cell volume regulation. Proc Natl Acad Sci USA 93: 12020-12025. Wang ZM, Katsurabayashi S, Rhee JS, Brodwick M, Akaike N. 2001. Substance P abolishes the facilitatory effect of ATP on spontaneous glycine release in neurons of the trigeminal nucleus pars caudalis. J Neurosci 21: 2983-2991. Watano T, Calvert JA, Vial C, Forsythe ID, Evans RJ. 2004. P2x receptor subtype-specific modulation of excitatory and inhibitory synaptic inputs in the brainstem. J Physiol 558: 745-757. Wieraszko A, Goldsmith G, Seyfried TN. 1989. Stimulation‐ dependent release of adenosine triphosphate from hippocampal slices. Brain Res 485: 244-250. Wirkner K, Ko¨falvi A, Fischer W, Gu¨nther A, Heike F, et al. 2005. Supersensitivity of P2X7 receptors in cerebrocortical

cell cultures after in vitro ischemia. J Neurochem 95: 14211437. Witting A, Walter L, Wacker J, Moller T, Stella N. 2004. P2X7 receptors control 2‐arachidonoylglycerol production by microglial cells. Proc Natl Acad Sci USA 101: 3214-3219. Xiang Z, Burnstock G. 2005. Expression of P2X receptors on rat microglial cells during early development. Glia 52: 119-126. Yao ST, Barden JA, Lawrence AJ. 2001. On the immunohistochemical distribution of ionotropic P2X receptors in the nucleus tractus solitarius of the rat. Neuroscience 108: 673-685. Yao SY, Ng AM, Muzyka WR, Griffiths M, Cass CE, et al. 1997. Molecular cloning and functional characterization of nitrobenzylthioinosine (NBMPR)‐sensitive (es) and NBMPR‐ insensitive (ei) equilibrative nucleoside transporter proteins (rENT1 and rENT2) from rat tissues. J Biol Chem 272: 28423-28430. Yegutkin GG, Henttinen T, Samburski SS, Spychala J, Jalkanen S. 2002. The evidence for two opposite, ATP‐generating and ATP‐consuming, extracellular pathways on endothelial and lymphoid cells. Biochem J 367: 121-128. Yoshioka K, Saitoh O, Nakata H. 2001. Heteromeric association creates a P2Y‐like adenosine receptor. Proc Natl Acad Sci USA 98: 7617-7622. Zhang JM, Wang HK, Ye CQ, Ge W, Chen Y, et al. 2003. ATP released by astrocytes mediates glutamatergic activity‐ dependent heterosynaptic suppression. Neuron 40: 971982. Zhang YX, Yamashita H, Ohshita T, Sawamoto N, Nakamura S. 1995. ATP increases extracellular dopamine level through stimulation of P2Y purinoceptors in the rat striatum. Brain Res 691: 205-212. Zhang YX, Yamashita H, Ohshita T, Sawamoto N, Nakamura S. 1996. ATP induces release of newly synthesized dopamine in the rat striatum. Neurochem Int 28: 395-400. Zimmermann H. 2000. Extracellular metabolism of ATP and other nucleotides. Naunyn Schmiedebergs Arch Pharmacol 362: 299-309. Zimmermann H. 2001. Ecto‐nucleotidases. In: Handbook of experimental pharmacology: Purinergic and pyrimidinergic signalling, Vol. 151/1. Abbracchio MP, Williams M editors. Berlin: Springer; pp. 209-251.

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Adenosine Neuromodulation and Neuroprotection

R. A. Cunha

1 1.1 1.2 1.3 1.4 1.5 1.6

Physiological Roles of Adenosine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 256 Adenosine as a Homeostatic Modulator . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 256 Pharmacology and Localization of Adenosine Receptors in the Brain . . . . . . . . . . . . . . . . . . . . . . . . . . . . 258 Neurotransmission and Neuromodulation – Adenosine as a Neuromodulator . . . . . . . . . . . . . . . . . . 260 Source of Endogenous Extracellular Adenosine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 263 Role of A1 Receptors in the Control of Synaptic Plasticity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 264 A2A Receptors and Modulation of Synaptic Plasticity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 265

2 Roles of Adenosine in Neurodegenerative Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 266 2.1 Therapeutic Opportunities to Manage Neurodegenerative Diseases Targeting the Adenosine Modulation System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 266 2.2 A1 Receptors as Hurls for the Development of Neuronal Dysfunction . . . . . . . . . . . . . . . . . . . . . . . . . . . . 267 2.3 Role of A2A Receptors in the Control of Neurodegeneration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 268 2.4 A2A Receptor Antagonists as Novel Anti‐Parkinsonian Drugs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 269 2.5 Role of A2A Receptors in Alzheimer’s Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 270 3

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Final Comments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 271

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Adenosine neuromodulation and neuroprotection

Abstract: The adenosine moiety fulfils an important intracellular homeostatic role since it is part of molecules that play key roles in defining the status of all cells, namely energy charge (ATP), redox status (NADH), and cell division (SAH/SAM). But the signaling role of the adenosine molecule itself is restricted to a paracrine role, signaling metabolic imbalance of cells within a tissue. Apart from this general role common to most tissues in mammals, adenosine fulfils a particular role as a neuromodulator in the nervous system. This involves a predominant inhibitory effect operated by adenosine A1 receptors, which results from a combined presynaptic inhibition of the release of excitatory neurotransmitters together with a postsynaptic action leading to neuronal hyperpolarization and an ability to depress plasticity by inhibition of NMDA receptors and voltage‐sensitive calcium channels. Overall, this A1 receptor‐mediated inhibition is aimed at decreasing the noise of excitatory transmission in brain circuits. Adenosine can also activate facilitatory A2A receptors, which only come into play at higher frequency of nerve stimulation and are directed at selectively shutting down A1 receptor inhibition in stimulated synapses to aid implementing changes in synaptic efficiency. Therefore, the combined role of A1 and A2A receptors is designed to increase salience of information in brain circuits. Apart from this main physiological role, adenosine also plays a relevant role in controlling the demise of damage in noxious brain conditions. In fact, the inhibitory A1 receptors are able to curtail brain damage. They play a role at the onset of brain damage and function as a hurl that needs to be overcome to allow the development of brain damage. In parallel, in chronic noxious brain conditions, A2A receptors contribute for brain damage, especially when insidious damage to synapses initiates neurodegeneration. Hence, A2A receptor antagonists are now being explored as novel neuroprotective strategies to interfere with the initial processes of neurodegenerative conditions, such as Parkinson’s and Alzheimer’s diseases. List of Abbreviations: CCPA, 2‐Chloro‐N6‐cyclopentyladenosine; CGS21680, 2‐[p‐(2‐carbonyl‐ethyl)‐ phenylethylamino]‐50 ‐N‐ethylcarboxamidoadenosine; Cl‐IB‐MECA, 1‐[2‐Chloro‐6‐[[(3‐iodophenyl)methyl] amino]‐9H‐purin‐9‐yl]‐1‐deoxy‐N‐methyl‐b‐D‐ribofuranuronamide; CP‐68,247, 8‐chloro‐4‐cyclohexyl‐ amino‐1‐(trifluoromethyl)[1,2,4]triazolo[4,3‐a] quinoxaline; CSC, 8‐(3‐chlorostyryl)caffeine; DPCPX, 1,3‐ dipropyl‐8‐cyclopentylxanthine; IB‐MECA, N6‐(3‐iodobenzyl)adenosine‐50 ‐N‐methyluronamide; KF17837, 1,3‐dipropyl‐8‐(3,4‐dimethoxystyryl)‐7‐methylxanthine; MRE3008F20, 5‐[[(4‐methoxyphenyl)amino] carbonyl]amino‐8‐ethyl‐2‐(2‐furyl)‐pyrazolo[4,3‐e]1,2,4‐triazolo[1,5‐c]pyrimidine; MRS1220, 9‐chloro‐ 2‐(2‐furanyl)‐5‐[(phenylacetyl)amino][1,2,4]‐triazolo[1,5‐c]quinazoline; MRS1334, 1,4‐Dihydro‐2‐methyl‐ 6‐phenyl‐4‐(phenylethynyl)‐3,5‐pyridinedicarboxylic acid 3‐ethyl‐5‐[(3‐nitrophenyl)methyl] ester; MRS1754, N‐(4‐cyano‐phenyl)‐2‐[4‐(2,6‐dioxo‐1,3‐dipropyl‐2,3,4,5,6,7‐hexahydro‐1H‐purin‐8‐yl)‐phenoxy]acetamide; R‐PIA, R‐N6‐(phenylisopropyl)‐adenosine; SCH58261, 5‐amino‐2‐(2‐furyl)‐7‐phenylethyl‐pyrazolo[4,3‐e]‐ 1,2,4‐triazolo[1,5‐c]pyrimidine; ZM241385, 4‐(2‐[7‐amino‐2‐[2‐furyl]‐[1,2,4]triazolo[2,3‐a]{1,3,5}triazin‐ 5‐yl‐amino]ethyl)phenol

1

Physiological Roles of Adenosine

1.1 Adenosine as a Homeostatic Modulator The viability of an organism ultimately depends on the ability of its cells to maintain a ‘‘constant’’ metabolism adapted to the needs of the organism (regulation) and in harmony with its environment. On the other hand, each cell should be able to change its metabolism according to modifications of the environment (control). This implies the need of devising signaling mechanisms (modulators) and devices (receptors and transducing systems), which allow sensing changes in the environment and implementing these changes inside cells to adequate their functioning. Since mammals are composed by cells organized in tissues, there is also the need to coordinate the response of each cell within a tissue (paracrine signaling) and of each tissue within the organism (endocrine signaling, which is aided by global control systems, such as the nervous or immune-inflammatory systems). One main paracrine signaling system is operated by adenosine (reviewed in Cunha, 2001). Adenosine is a purine nucleoside which normally occurs in concentrations of tenths of nanomolar within virtually all

Adenosine neuromodulation and neuroprotection

11

eukaryotic cells. However, considerably higher concentrations of chemical derivatives of this nucleoside are present within cells (> Figure 11‐1): for instance, it is present in the form of S‐adenosylhomocysteine (SAH) and S‐adenosylmethionine (SAM), nicotanimide and adenine nucleotides (NADþ and NADPþ) and adenine nucleotides (AMP, ADP, and ATP). In this sense, the adenosine moiety is directly involved in the three fundamental systems that regulate the functioning of eukaryotic cells: cell cycle (SAH/SAM), redox potential (NADH/NADþ), and energy charge (ATP, ADP, and AMP). However, the signaling role of the adenosine molecule as such is reserved to a particular situation of paracrine signaling of energetic modification with a tissue. . Figure 11‐1 Adenosine derivatives are directly involved in the three fundamental systems that regulate the functioning of eukaryotic cells. In fact, S‐adenosylhomocysteine (SAH) and S‐adenosylmethionine (SAM) control cell cycle, nicotanimide and adenine nucleotides (NADþ and NADPþ) define the redox state and adenine nucleotides (AMP, ADP, and ATP) define the energy status of cells. However, the signaling role of the adenosine molecule as such is reserved to a particular situation of paracrine signaling of energetic modification with a tissue

In a situation of energetic (dynamic) ‘‘equilibrium’’ between the generation and requirement of energy (ATP for the sake of simplicity), the concentration of this nucleotide is kept constant at a value of 3–10mM in different eukaryotic cells. In a situation of metabolic imbalance in a particular cell within a tissue, either because there is a shortage of oxidative potential or because there is an increased workload, the cell becomes metabolically stressed. This condition needs to be signaled to all neighboring cells in the tissue to enable them to rapidly adapt their metabolism to new environmental, potentially stressful conditions. Given the importance of the maintenance of tissue homeostasis, this signaling ought to be rapid, while the particular cell that first sensed the stressful change of environment is still beginning to become energetically imbalanced. This is possible, thanks to the difference in the extracellular concentration of ATP (circa 5mM) and of adenosine (circa 50nM). Hence, the concentration of adenosine will rise 100,000‐fold with a change of 1% of the concentration of ATP. This change in intracellular adenosine is converted to a gradient of extracellular adenosine around this cell since all eukaryotic cells are equipped with bidirectional nonconcentrative nucleoside (adenosine) transporters. These increased levels of intracellular adenosine can then activate the most abundant and ubiquitous adenosine receptor subtype, the A1 receptor, which has the ability of decreasing metabolic flow in eukaryotic cells (reviewed in Cunha, 2001) (> Figure 11‐2).

257

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. Figure 11‐2 Adenosine fulfils a paracrine role signalling situations of metabolic imbalance within tissues. When a stressful event occurs in a given cell (left part of the figure), ATP is used to attempt restoring the functioning of this cell. As a minor fraction of ATP is used, there is a massive formation of adenosine because of the near 250,000‐fold difference in their concentrations. This adenosine outflows from this cell through bidirectional and nonconcentrative nucleoside transporters (T). This adenosine gradient acts through adenosine A1 receptors (A1R) in neighboring cells where it decreases the metabolic rate allowing these cells to cope better with potential stressful stimuli that may reach them

This decrease of metabolic rate allows cells to cope better with stressful conditions and also increase their sensitivity to recruitment signals (by decreasing noise). This generic system of tissue paracrine signaling of metabolic imbalance has been exploited in several tissues as a signaling mechanism to coordinate tissue activity. This can be exemplified by the role of adenosine in the control of pacemaking in the heart (Shryock and Belardinelli, 1997), in the control of tubuloglomerular filtration rate in the kidney (Osswald et al., 1996), in the control of postprandial vasodilatation in the liver (Lautt, 1996), in the control of immune‐inflammatory reactivity to limit collateral damage (Sitkovsky et al., 2004), or in the control of the activity of neuronal circuits in the brain (reviewed in Dunwiddie and Masino, 2001). This homeostatic inhibitory effect of adenosine in the central nervous system is expected to be most relevant since the brain is equipped with a density of adenosine A1 receptors considerably greater than that of other tissues (reviewed in Fredholm et al., 2005). This is best illustrated by the fact that the consumption of caffeine, which only known mechanism of action at nontoxic concentrations is the antagonism of adenosine receptors (Fredholm et al., 1999), causes effects involving the modification of brain functioning (reviewed in Fredholm et al., 1999).

1.2 Pharmacology and Localization of Adenosine Receptors in the Brain Adenosine can act through the activation of four different adenosine receptors (A1, A2A, A2B, and A3). These receptors constitute the subfamily of P1 receptors to distinguish them from P2 receptors which are activated by adenosine‐50 ‐ and uridine‐50 ‐di‐ and tri‐phosphate (Ralevic and Burnstock, 1998). Adenosine receptors are G‐protein coupled receptors (see > Table 11‐1), which have limited sequence homology between each other in the different species where they were cloned (reviewed in Fredholm et al., 2005). The A1 receptor is the most abundantly expressed and its primary sequence is most conserved between species in the family. A1 Receptors couple to Gi/o proteins but their ability to inhibit neuronal activity may not only depend on their ability to inhibit cAMP accumulation and activate phospholipase C activity, but also to signal through b,g subunits of G proteins (Fredholm et al., 2005). A1 Receptors are widespread and abundantly expressed in the brain with a pre‐ and postsynaptic localization and with high levels in the cerebral cortex,

11

Adenosine neuromodulation and neuroprotection

. Table 11‐1 Represents the pharmacological properties of the most important adenosinergic ligands at the rat (r) and the human (h) receptors. Values are bold in columns of those receptors for which the ligand has the selectivity Agonist Adenosine

rA1

hA1 54

73 Inosine CCPA

8.1 0.4

R‐PIA CGS21680 IB‐MECA Cl‐IB‐MECA Antagonist DPCPX

3,100 54

rA1 0.73 0.3 0.6

SCH58261 caffeine ZM241385 CSC KF17387 CP‐68,247 MRS1334 MRS1220 MRE3008F20 MRS1754

rA2A 150

6,700 6.4 0.8 2.0 290 3.7 120 hA1 1.6 3.9

12,000 540

138 1,493 438 87 >100,000 305

3,900

22 56

hA2B 11,300

5,100

rA3

hA3 56

6,500

2,300 860 27

42 16 67

11,200a 88,800a

2,500 2,100

1.1 0.3

hA2A

340 682

130

50

4000

0.6

5,000b

10,000

1.2 8,100

2,400 1.7

rA2B

17,000

hA2B

13,000 31

0.67 61 17 41,561 >100,000 52 140 500 610

2,100 2.0

rA3

1.2 11

rA2A

1,100 400 17

rA2B

81

290 740 20,000

hA2A 960

hA3

190,000

80,000 270

3,850 >1,000

2.7 0.7 0.29 570

Ref. 1 2 1 3 4 4 4 11 4 4 Ref. 3 5 7 5 7 6 6 7 7 7 7 6 6 8 9 10

References: (1) Rank order of potency (EC50) values, in Chinese hamster ovary (CHO) cells transfected with human adenosine receptors (Fredholm et al., 2001b); (2) Rank order of potency (EC50) values, in Chinese hamster ovary (CHO) cells transfected with rat adenosine receptors (Yaar et al., 2004); (3) Binding affinities (Ki values) in rat brain cortical membranes and membranes of CHO cells expressing the human A1 receptor (Wittendorp et al., 2004); (4) Comparison of affinity of agonists at rat and human adenosine receptor subtypes (Ki values in nM). aEC50‐values (nM) for the agonist‐mediated stimulation of adenylyl cyclase activity in a membrane preparation (Klotz, 2000); (5) Binding affinity (Ki values) of antagonists at adenosine receptor subtypes. bInhibition of cAMP accummulation (Klotz, 2000); (6) Potency (KD values) of caffeine at rat and human adenosine receptor subtypes (Fredholm et al., 1999); (7) Potency of some adenosine receptor antagonists to displace CGS21680 from adenosine A2A receptors CHA from adenosine A1 receptors in rat striatum. Results are given as Ki values in nM (Fredholm and Lindstro¨m, 1999); (8) Binding affinity at human adenosine receptor subtypes in CHO cells (Ki values), (Varani et al., 2000). Not a useful ligand at rat receptors; (9) Binding affinity at human adenosine receptor subtypes in transfected HEK 293 cells (Ki values), (Kim et al., 2000); (10) Binding affinity at rat adenosine receptor subtypes in transfected HEK 293 cells (Ki values), (Kim et al., 2000); (11) Given are IC50 values, radioligands [3H]CHA (A1), [3H]NECA (A2A) (Hutchison

hippocampus, cerebellum, thalamus, brain stem, and spinal cord of rodents and Humans (see Fredholm et al., 2005). Adenosine A2A receptors are expressed at comparatively lower levels in the nervous system, with the exception of the basal ganglia, where they are expressed at exceptionally high levels in GABAergic medium spiny neurons (Svenningsson et al., 1999). Activation of A2A receptors has effects largely opposite

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to these of A1 receptors. Although they are mostly viewed as signaling through GS proteins, several neuronal effects of A2A receptors apparently involve other transducing systems (Fredholm et al., 2005), which still await to be clarified. Two other adenosine receptors have been cloned and pharmacologically characterized, namely A2B and A3 receptors. However, they have an expression lower than that of A1 and A2A receptors in the brain and a limited number of studies have so far explored their eventual role in controlling brain function (Fredholm et al., 2005). Adenosine is the primary full agonist at all four adenosine receptor subtypes. There are several adenosine receptor agonists which are mainly derivatives of adenosine with N6 and C2 substitutions of the adenine base and C5 substitutions of the ribose moiety (Fredholm et al., 2001). As illustrated in > Table 11‐1, these agonists effectively distinguish between A1, A2A, and A3 receptor mediated effects in in vitro experiments, whereas selective A2B receptor agonists await widespread acceptance, especially because this receptor has lower affinity for adenosine and for most agonists. The situation is less fortunate in vivo as the pharmacokinetics of these compounds have not been studied extensively. To be considered selective, the potency of a considered ligand between different adenosine receptors should ideally differ by at least two orders of magnitude. However, this is not always the case with adenosine receptor agonists, thus limiting their use for studying specific receptor function in vivo (Fredholm et al., 2001). There are no known endogenously produced adenosine receptor antagonists in mammals. However, there are several selective antagonists for the different adenosine receptors (see > Table 11‐1). The initial generation was constituted by derivatives of caffeine, the mostly widely consumed psychoactive drug (Fredholm et al., 1999). However, several different classes of compounds have been developed as pharmacological tools with appreciable selectivity for the different adenosine receptors (see > Table 11‐1). As for agonists, there are better and widespread used antagonists to manipulate A1 and A2A receptors. The A3 receptors display considerable difference in their profile of antagonist sensitivity between different species. Finally, the novel A2B receptor antagonists still await widespread validation. Hence, the combined density of the different adenosine receptors together with the availability of adequate pharmacological tools enables to consider adenosine neuromodulation in brain circuits as being mainly processed by A1 and A2A receptors, albeit it should be kept in mind that the two other receptors (A2B and A3) may also play so far unravelled roles.

1.3 Neurotransmission and Neuromodulation – Adenosine as a Neuromodulator From the conceptual point of view, the functioning of the brain is based on its ability to process in parallel, compare and select the relevant signals, which are coded in the form of frequency and simultaneity of firing of action potentials in neuronal circuits (see Rieke et al., 1999). Each neuron takes an instantaneous all‐or‐ none decision of firing an action potential (as fast as every 2 milliseconds) based on the computation of the excitatory and inhibitory signals received from all afferent neurons (each neuron contacts an average of 10,000 other neurons). For this purpose, the majority (near 90%) of neurons in the brain uses two main neurotransmitters: glutamate, the main excitatory neurotransmitter and GABA, the main inhibitory neurotransmitter. Therefore, the pharmacological manipulation of either of these two main transmitter systems with therapeutic goals is likely to cause widespread effects in nearly all brain circuits. This obviously limits the therapeutic targeting of the glutamatergic and GABAergic systems to situations where there are global changes of brain functioning. In contrast, their use to correct dysfunction of a particular circuit is expected to cause side effects in numerous other circuits functioning normally. The complexicity of functioning of brain circuits is not only due to the number of wired neurons and to their speed of functioning but also the numerous modulation systems designed to fine‐tune neuronal circuits to enable them to function in an optimal homeostatic manner. The main characteristic of these neuromodulation systems is that they do not directly vehicle information but rather control the efficiency of functioning of the communication systems operated by neurotransmitters (see Katz, 1999). From the therapeutic point of view, these modulation systems represent attractive targets since several of them display

Adenosine neuromodulation and neuroprotection

11

regional and event circuit selectivity and only fine‐tune rather than directly trigger dramatic (all‐or‐none) responses, and consequently are expected to trigger less secondary side effects. Adenosine is one important neuromodulation systems in the brain. It mainly acts in excitatory synapses as detailed by numerous studies in vitro and in vivo in different animal species as well as in Humans (reviewed in Dunwiddie and Masino, 2001). The administration of adenosine or of nondegradable chemical derivatives of adenosine causes a profound depression of excitatory synaptic transmission (Dunwiddie and Masino, 2001). This effect depends on the activation of inhibitory A1 receptors, since it is prevented by xanthine derivatives that selectively antagonise A1 receptors, such as 1,3‐dipropyl‐8‐ cyclopentylxanthine (DPCPX) (Dunwiddie and Masino, 2001). Most importantly, it is found that the addition of a concentration (or dose) of an A1 receptor antagonist facilitates excitatory synaptic transmission (Dunwiddie and Masino, 2001). This shows that this adenosine inhibitory modulation system is continuously and constitutively active in brain circuits (Dunwiddie and Masino, 2001), which means that extracellular adenosine is continuously produced during the functioning of brain circuits (> Figure 11‐3). This is the reason why the consumption of caffeine (an adenosine receptor antagonist) causes evident effects due to the modification of functioning of brain circuits (Fredholm et al., 1999). . Figure 11‐3 Adenosine profoundly inhibits synaptic transmission and tonically inhibits synaptic transmission in functioning brain circuits. This representative experiment was carried out in rat hippocampal slices (drawn on the left of the figure). Following recovery after dissection, the slices were stimulated by placing a stimulation electrode over the afferent Schaffer fibers and recording were carried out either in the cell body layer of the stratum pyramidale (SP, allowing the recording of population spikes) or in the synaptic region of the stratum radiatum (SR, allowing the recording of excitatory postsynaptic potencials, fEPSPs), as illustrated in the left side of the figure. These fEPSPs were recorded over time and the effect of different drugs, applied through the superfusion solution, was tested, when indicated by the bars above the figure in the right panel of the figure. Adenosine (Ado, 10 mM) and its closest nonmetabolizable analog, 2‐chloroadenosine (CADO, 10 mM) blocked synaptic transmission in a reversible manner, whereas they were virtually devoid of effects in the presence of a supramaximal concentration of the selective A1 receptor antagonist, DPCPX (50 nM). Note that DPCPX caused a 15% facilitation of synaptic transmission indicating that endogenous adenosine is tonically depressing synaptic transmission. This experiment also shows that the main effect of adenosine is an A1 receptor‐mediated inhibition of synaptic transmission and no other effects of adenosine are observable upon blockade of A1 receptors

The molecular mechanisms underlying this predominant inhibitory effect operated by activation of A1 receptors are displayed in > Figure 11‐4 and have been explored by different groups (reviewed in Fredholm et al., 2005). It was concluded that this inhibition mainly results from the effects operated by A1 receptors located in the active zone of presynaptic nerve terminals, which inhibit the evoked release of glutamate by controlling calcium entry and the sensitivity to calcium of the fast exocytotic release machinery (reviewed

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. Figure 11‐4 The inhibition of synaptic transmission and neuronal excitability operated by A1 receptors involves pre‐, post‐, and nonsynaptic effects. The inhibition of synaptic transmission mainly results from the action of presynaptic A1 receptors (A1) located in the active zone of excitatory nerve terminals, which inhibit the evoked vesicular release of glutamate (Glu) by a combined inhibition of calcium entry and decreased sensitivity of the release apparatus to calcium. The inhibition of synaptic plasticity mainly results from the action of postsynaptic A1 receptors, located in the postsynaptic density, which can decrease the entry of calcium through the inhibition of voltage‐sensitive calcium channels (VSCC) mainly of the N‐type as well as by inhibition of NMDA receptors. Finally, the inhibition of neuronal excitability mainly results from the action of nonsynaptic A1 receptors, which controls potassium conductances leading to a neuronal hyperpolarization. This last effect is mainly responsible for the ability of A1 receptors to control convulsive behavior

in Fredholm et al., 2005). The efficiency of this inhibitory system is reenforced by the presence of A1 receptors located at the postsynaptic density, which control the entry of calcium through voltage‐ sensitive calcium channels and through NMDA receptors. Postsynaptic A1 receptors located in dendrites also activate potassium conductance, which inhibit neuronal firing and contribute for the inhibition of synaptic transmission (reviewed in Fredholm et al., 2005). Combined neurochemical and electrophysiological studies allowed concluding that the extracellular levels of adenosine increase with increasing frequency of firing of excitatory fibers, which is expected to increase the inhibitory effect operated by A1 receptors (Dunwiddie and Masino, 2001). Based on the general scenario whereby A1 receptors act as a homeostatic inhibitory signal upon increased workload in different peripheral tissues, it was hypothesized that A1 receptors would also act as a feedback inhibitory system to restraint excessive excitatory transmission. This hypothesis proposed that an increased activity of an excitatory synapse would imply a greater consumption of ATP to restore ionic balance, which would lead to a greater production of adenosine; these increased levels of extracellular adenosine acting through A1 receptors would gradually increase the feedback inhibition of this synapse counteracting the excessive activation and preventing synaptic damage (reviewed in Cunha, 2001; Dunwiddie and Masino, 2001).

Adenosine neuromodulation and neuroprotection

11

1.4 Source of Endogenous Extracellular Adenosine One topic of fundamental importance to understand adenosine neuromodulation, which is still ill defined, is the mechanisms by which adenosine is released into the extracellular medium. Adenosine can appear in the extracellular milieu mainly through two different mechanisms (> Figure 11‐5): (1) the release of . Figure 11‐5 Extracellular adenosine in brain tissue can result from the extracellular catabolism of released ATP or from a release of adenosine as such, both of which can be originated in nerve terminals or dendritic regions, from activated astrocytes and even from microglia. Nerve terminals can release both ATP and adenosine, the former from synaptic vesicles and the later through nonconcentrative bidirectional nucleoside transporters. Activated dendrites can also release both ATP and adenosine by mechanisms still to be resolved. Likewise, activated astrocytes can also release ATP in a vesicular manner as well as adenosine by unresolved mechanisms. Microglia can also release both ATP and adenosine (unpublished results) by unresolved mechanisms. All cell types in the brain as well as all subsynaptic compartments are equipped with different types of ectonucleotidases able to convert ATP into adenosine (represented by the grey circles). These ectonucleotidases have a notable efficiency, which involves channelling mechanisms, allowing the formation of adenosine from extracellular ATP in less than 20 milliseconds, making it difficult to distinguish the metabolic source of extracellular adenosine

adenosine as such through nucleoside transporters (Geiger and Fyda, 1991) following an increase of the intracellular levels of adenosine or a reversal of the sodium gradient; (2) the extracellular formation of adenosine through the ectonucleotidase pathway on release of adenine nucleotides, especially ATP (reviewed in Dunwiddie and Masino, 2001; Latini and Pedata, 2001). Release of adenosine after its intracellular formation has been clearly shown following hypoxia or metabolic poisoning and following field stimulation of brain slices (see Cunha, 2001; Dunwiddie and Masino, 2001; Latini and Pedata, 2001). Under such conditions, there is an imbalance between energy supply and demand, leading to a net hydrolysis of intracellular ATP. This intracellular ATP is then converted into adenosine, leading to a dramatic increase in the intracellular concentration of adenosine. The presence of nonconcentrative bidirectional nucleoside transporters will then force extracellular adenosine levels to rise in parallel with intracellular adenosine. However, the release of adenosine through nucleoside transporters appears to be at odds with the ability of inhibitors of nucleoside transport to potentiate the effects of endogenous adenosine (reviewed in Latini and Pedata, 2001; Cunha, 2001), which indicates that the role of nucleoside transporters

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is mostly to clear up rather than to mediate adenosine release, at least in nonstressful situations. The importance of the type 1 equilibrative nucleoside transporter (ENT‐1) in regulating extracellular adenosine levels has recently also been demonstrated in knockout mice lacking this protein (Choi et al., 2004). ENT‐1 knockout mice exhibit a reduced adenosine tone reflected by a decreased adenosine‐mediated inhibition of glutamate excitatory postsynaptic currents. This result is compatible with ENTs being involved in releasing adenosine, so the issue remains unsettled. However, the recent evidence gathered by Boison’s group on the key role of adenosine kinase in controlling the extracellular levels of adenosine in brain tissue (Boison, 2006) clearly argues in favor of a major role of nucleoside transporters in clearing up rather than in releasing adenosine in brain tissue. The alternative hypothesis is to consider that adenosine is formed by extracellular catabolism of released adenine nucleotides, mainly ATP, through the ectonucleotidase pathway (see Zimmermann, 2000) (> Figure 11‐5). ATP can be released from vesicles from nerve terminals (see Cunha, 2001), but also from astrocytes (reviewed in Halassa et al., 2007). There are some direct evidence supporting the view that the extracellular catabolism of ATP acts as source of adenosine, but it is still unclear if this ATP is mainly released from nerve terminals, from astrocytes or from either structures according to the pattern of recruitment of neuronal circuits (reviewed in Fields and Burnstock, 2006; Cunha, 2007). Although it still needs to be determined what is the source and mechanism of adenosine release, it is nevertheless clear from the elegant work of Dale’s group that ectonucleotidases can play a crucial role in controlling the buildup of adenosine as a function of neuronal activity (Dale, 2002), although this finding could not be reproduced using different preparations where no clear‐cut separation of neurotransmission and control of tissue viability could be achieved (see Cunha, 2001; Dunwiddie and Masino, 2001; Latini and Pedata, 2001).

1.5 Role of A1 Receptors in the Control of Synaptic Plasticity One of the most remarkable features of the brain is the ability of its neuronal circuits to modify their intrinsic properties of efficiency of transmission as a function of their recruitment (Lynch, 2004). This property is now considered to represent the neurophysiological basis of implementation of memory traits resulting from learning experiences (Lynch, 2004). At the synaptic level, this property can be measured as an increased efficiency of synaptic transmission upon either increasing the firing frequency of particular afferent fibers or promoting the simultaneity of arrival of signals from different incoming fibers, the two main coding strategies in brain circuits (Lynch, 2004). In fact, particular firing frequencies of glutamatergic synapses (or particular coincidence of firing of different synapses onto the same neuron) can trigger an increase in the efficiency of functioning of these particular synapses, which can persist long after the triggering event has occurred. These phenomena are called long‐term potentiation. For instance, in hippocampal synapses, a limbic cortical region involved in spatial learning abilities amongst others (Squire et al., 2004), a single episode of high frequency stimulation (100 stimuli applied for 1 second) leads to an increase efficiency of the stimulated synapses that can last weeks (see Lynch, 2004). This means that the response to a particular electrical stimulus after the high frequency episode is greater than the response to the same stimulus before this frequency‐dependent stimulation episode. The implementation of this increased efficiency of synaptic transmission mainly requires the recruitment of NMDA receptors, which do not participate significantly to synaptic transmission at basal (low frequency, i.e., Section 3), as well as receptor tyrosine kinases (> Section 4), regulate AMPAR function in several brain regions. > Table 12-1 summarizes the reviewed literature on the regulation of AMPAR by metabotropic receptors and receptor tyrosine kinases.

2

Levels of AMPA Receptor Regulation

2.1

AMPA Receptor Structure

AMPAR are tetrameric assemblies of the GluR1–GluR4 subunits, which form either homomeric or heteromeric receptors, with distinct functional properties, depending on the subunit composition and . Table 12-1 Modulation of AMPA receptors by metabotropic receptors and receptor tyrosine kinases Receptor ligand Glutamate

Receptor type Group I mGluRs Group I mGluRs Group I/II mGluRs Group I/II mGluRs Group I/II mGluRs Group I mGluRs

mGluR5 mGluR5 Group I/II mGluRs mGluR1

mGluR1

Cell type/ tissue Spinal cord motoneurons Rat spinal dorsal horn neurons Rat visual cortex NTS Cerebellar Purkinje neurons Cultured hippocampal neurons

Hippocampal slices Hippocampal slices Cultured chick Purkinje neurons Cultured Purkinje neurons Cerebellar slices

Synaptic effect ↑ AMPA‐induced membrane depolarization ↑ AMPA‐induced currents ↑ AMPA‐evoked [Ca2þ]i rise ↑ AMPA induced membrane depolarization ↑ AMPA‐induced currents

Reference (Ugolini et al., 1997, 1999) (Bleakman et al., 1992; Cerne and Randic 1992) (Wang and Daw 1996; Wang et al., 1998) (Glaum and Miller 1993)

↑ AMPA induced membrane depolarization

(Glaum et al., 1992)

↓ Synapses with AMPA receptor clusters (GluR1, GluR2) ↑ GluR1 at the cell body ↓ AMPA receptor‐mediated mEPSCs ↑ LTD ↓ Surface GluR1 ↑ Depotentiation of previously established LTP ↓ AMPA induced membrane depolarization

(Snyder et al., 2001)

(Xiao et al., 2001; Zho et al., 2002; Huang et al., 2004) (Zho et al., 2002) (Mori‐Okamoto et al., 1993)

↑ LTD

(Shigemoto et al., 1994)

↑ LTD

(Aiba et al., 1994; Conquet et al., 1994; Hartell 1994)

Regulation of AMPA receptors by metabotropic receptors and receptor tyrosine kinases

12

. Table 12-1 (continued) Receptor ligand Dopamine

Receptor type D1 receptors

Cell type/ tissue Dorsal striatum

D2 receptors

Dorsal striatum

D1 receptors

Nucleus accumbens

D1 receptors

Prefrontal cortex

D2 receptors

Prefrontal cortex Retina

D1 receptors

Serotonin

Adenosine Acetylcholine

Noradrenaline

5‐HT2 serotonin receptors

Spinal cord dorsal horn neurons (young animals)

5‐HT2 serotonin receptors 5‐HT1A serotonin receptors

Trigeminal motor neurons Prefrontal cortical pyramidal neurons Aplysia siphon motor neurons

5‐HT (Ap5‐HTB2) serotonin receptors A2 adenosine receptors Muscarinic receptors b‐adrenergic receptors

Hippocampal CA1 region Hippocampal CA1 pyramidal neurons Hippocampal CA1 neurons

Synaptic effect ↑ AMPAR EPSPs ?Rundown of iontophoretic AMPAR currents ↑ LTP ↓ AMPAR EPSPs ? LTP

Reference (Umemiya and Raymond 1997; Lin et al., 2003) (Yan et al., 1999) (Calabresi et al., 2000; Kerr and Wickens 2001) (Cepeda et al., 1993; Levine et al., 1996) (Calabresi et al., 1997) ↑ GluR1 phosphorylation at (Chao et al., 2002b) (Chao Ser845 et al., 2002a) ↑ GluR1 surface expression ↑ AMPAR EPSCs (Gonzalez‐Islas and Hablitz 2003) ↑ GluR1 surface (Wolf et al., 2003; Swayze expression et al., 2004) ↑ LTP (Gurden et al., 2000) ↓ AMPAR‐mediated (Tseng and O’Donnell 2004) responses ↑ [Ca2þ]i response to (Gomes et al., 2004) kainate ↑ GluR4 phosphorylation at Ser842 ↑ GluR4 surface expression ↑ AMPAR EPSCs (Li et al., 1999) Unsilencing of glutamatergic synapses between sensory afferents and spinal cord dorsal horn neurons ↑ Iontophoretic AMPA (Trueblood et al., 1996) responses ↓ AMPA‐evoked currents ↓ GluR1 phosphorylation at Ser831

(Cai et al., 2002b)

↑ AMPAR postsyaptic potentials

(Chitwood et al., 2001)

↑ AMPAR EPSPs

(Kessey and Mogul 1997)

↑ AMPAR EPSPs

(Auerbach and Segal 1996)

↑ GluR1 phosphorylation at Ser845

(Vanhoose and Winder 2003)

279

280

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Regulation of AMPA receptors by metabotropic receptors and receptor tyrosine kinases

. Table 12-1 (continued) Receptor ligand Opioids Somatostatin

Vasopressin

Oxytocin

Substance P

NKA

NKB Brain natriuretic peptide Insulin

Receptor Cell type/ type tissue Synaptic effect k‐ and m‐opioid Dorsal horn ↓ AMPA‐evoked currents receptors neurons ↓ AMPAR‐mediated Somatostatin ‐Mediobasal responses sst2 receptors hypothalamic slices ‐ Mediobasal hypothalamic cultures ↓ AMPA‐evoked currents Vasopressin Vasopressin V1a receptors neurons in the supraoptic nucleus ↑ AMPA‐evoked currents Oxytocin Oxytocin receptors neurons in the supraoptic nucleus Tachykinin NK1 Dorsal and ↑ AMPA‐evoked currents receptors ventral horn neurons Tachykinin NK2 Dorsal and ↑↓ AMPA‐evoked currents receptors ventral horn neurons Tachykinin NK3 Dorsal horn ↑ AMPA‐evoked currents receptors neurons B‐type Nucleus of the ↑ AMPA‐evoked currents natriuretic tractus peptide solitarius receptors ↑ GluR1 and GluR2 Insulin Cultured internalisation receptors hippocampal neurons Insulin receptors Insulin receptors

IGF‐1

IGF‐1 receptor

BDNF

Trk B receptors Trk B receptors

Hippocampal CA1 neurons Mossy fibers to CA3 pyramidal cell synapses Cultured cerebellar Purkinje neurons Cultured neocortical neurons Nucleus tractus solitarius

Reference (Yoshimura and Jessell 1990; Kolaj et al., 1995) (Peineau et al., 2003)

(Hirasawa et al., 2003)

(Hirasawa et al., 2003)

(Rusin et al., 1992; Rusin et al., 1993; Cumberbatch et al., 1995) (Rusin et al., 1992; Rusin et al., 1993; Chizh et al., 1995; Cumberbatch et al., 1995) (Cumberbatch et al., 1995) (Glaum and Miller 1993)

(Beattie et al., 2000; Lin et al., 2000; Man et al., 2000; Zhou et al., 2001; Ahmadian et al., 2004) (Ahmadian et al., 2004)

↓ AMPAR‐mediated currents ↑ LTD

(Huang et al., 2003)

↓ AMPAR‐evoked currents

(Wang and Linden 2000)

↑ AMPA receptor delivery to the plasma membrane

(Narisawa‐Saito et al., 2002)

↓ AMPAR‐evoked currents

(Balkowiec et al., 2000)

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12

stoichiometry, and differential expression in vivo throughout the central nervous system. In addition to the diversity provided by the different genes and different combinations of subunits, alternative splicing and RNA editing introduce additional diversity. The four AMPAR subunits have an alternatively spliced region known as ‘‘flip/flop,’’ which influences receptor desensitization (Sommer et al., 1990); the flip version of the receptors desensitizes four times slower than the flop version (Mosbacher et al., 1994). Alternative splicing also occurs at the regions encoding the C‐terminus of GluR2 and GluR4, generating short and long forms of these two AMPAR subunits (Gallo et al., 1992; Kohler et al., 1994). RNA editing also has an important impact on receptor function (Seeburg, 1996). RNA editing is mediated by ADARS (adenosine deaminases acting on RNA), which recognize a complementary intronic sequence, and change one base to alter the amino acid sequence (Seeburg and Hartner, 2003). RNA editing affects the ion channel permeability of AMPAR containing the GluR2 subunit. The editing of GluR2 at the Q/R site gives rise to an arginine residue in the ion pore forming region of the protein, which makes GluR2‐containing AMPAR impermeable to calcium and nonrectifying (Sommer et al., 1991). RNA editing at the R/G site, before the flip/flop region, affects AMPAR recovery from desensitization (Lomeli et al., 1994). AMPAR are fully activated and desensitized by AMPA, quisqualate, and glutamate; kainate produces a partial but less desensitizing activation of these receptors. The topology of each AMPAR subunit is well established, and the accepted model predicts the existence of a large N‐terminal extracellular domain, three transmembrane segments, a membrane reentrant pore loop, an extracellular loop between transmembrane domains M3 and M4, and an intracellular C‐terminal segment (> Figure 12-1). The membrane proximal part of the N‐terminus of the protein, together with part of the extracellular loop that connects transmembrane domains M3 and M4, forms the ligand‐binding region of AMPAR (S1S2, see > Figure 12-1). Crystal structures have provided high‐resolution pictures of the structure of the glutamate‐binding domain, and have led to suggestions as to how ligand‐binding controls channel gating and desensitization (Armstrong et al., 1998; for a review see, Oswald, 2004). The ligand binds in a cleft between the two lobes of the S1S2 domain and makes contacts with both lobes. The two lobes of the protein close upon binding to the agonist, and the partial agonist kainate induces partial closure of the cleft. Most cues for the structure of the ion pore of glutamate receptors have been inferred in comparison with Kþ channels. The homology modeling and experimental evidence collected so far are consistent with the pore‐forming domains in GluR and Kþ channels with a similar structure but inverted membrane topology (Oswald, 2004). Interestingly, some data suggest that, in addition to the membrane reentrant pore M2, the transmembrane domain M3 also plays a role in gating (Jones et al., 2002; Sobolevsky et al., 2003). The C‐terminal portion of AMPAR subunits is intracellular and important in terms of regulating the function and localization of the receptors. Although the extracellular and transmembrane regions of AMPAR subunits are very similar, their intracellular cytoplasmic tails are distinct and can be divided into two groups. GluR1, GluR4, and the alternatively spliced long form of GluR2, GluR2long, are homologous and have long cytoplasmic tails, whereas GluR2short, the predominant splice version of GluR2, GluR3, and GluR4c, the cerebellum‐expressed splice variant of GluR4, have short, homologous, cytoplasmic C‐termini (> Figure 12-1). Each subunit interacts with specific cytoplasmic proteins through the C‐terminal region (see later, Bredt and Nicoll, 2003). Moreover, this region of the AMPAR harbors phosphorylation sites and their phosphorylation directly regulates the activity of the receptors (Carvalho et al., 2000) and plays a role in governing their cellular traffic (Bredt and Nicoll, 2003; Gomes et al., 2003).

2.2

AMPA Receptors and Synaptic Plasticity

Activity‐dependent changes in synaptic activity are ubiquitous in various brain regions, and are thought to underlie the processes of learning and memory formation. The most prominent examples of long‐lasting activity‐dependent changes in synaptic strength are long‐term potentiation (LTP) and long‐term depression (LTD). Their mechanisms have been extensively studied in the last 10 years (for a review see, Malenka and Bear, 2004). Their mechanisms vary depending on the synapses where they occur, but it is now clear that a major mechanism for the expression of LTP involves an increase in the number of AMPAR in the plasma membrane at synapses, via activity‐dependent changes in AMPAR traffic (Malinow and Malenka, 2002;

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. Figure 12-1 AMPAR exist at postsynaptic sites on spines and extrasynaptically (a). Topology of AMPAR subunits (b) and comparison of their C‐terminal region (c). Identified phosphorylation sites are underlined

Song and Huganir, 2002; Bredt and Nicoll, 2003) and modification of the biophysical properties of AMPAR as a consequence of receptor phosphorylation (Benke et al., 1998; Soderling and Derkach, 2000). On the other hand, LTD in the hippocampus is associated with a selective dephosphorylation of the protein kinase A (PKA) phosphorylation site on GluR1 [see later; (Lee et al., 1998, 2000, 2003)], accompanied by the physical loss of AMPAR from the synapse, as a consequence of rapid receptor internalization through a clathrin‐ and dynamin‐dependent mechanism (Beattie et al., 2000; Ehlers, 2000; Lin et al., 2000).

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12

AMPA Receptor Phosphorylation

Phosphorylation of ligand‐gated ion channels is crucial in the regulation of their function, and plays an important role in the mechanisms of synaptic plasticity (Soderling and Derkach, 2000). Phosphorylation of AMPAR regulates their activity and affects their interaction with intracellular proteins as well as their expression at the cell surface and at the synapse (Malinow and Malenka, 2002; Song and Huganir, 2002; Bredt and Nicoll, 2003). Growing literature supports a role for AMPAR trafficking, which is largely conditioned by AMPAR phosphorylation, in activity‐dependent changes in synaptic function (Malinow and Malenka, 2002). Initial studies demonstrated the importance of protein kinase activity in the regulation of AMPAR function by changing protein kinase activity in neurons, using appropriate drugs (for a review see, Carvalho et al., 2000) or by intracellular perfusion with inhibitor peptides (Greengard et al., 1991) or with constitutively active kinases (McGlade‐McCulloh et al., 1993; Wang et al., 1994), and looking for the resulting effects on AMPAR function. Later biochemical analysis of AMPAR phosphorylation found evidence for the direct phosphorylation of AMPAR subunits by intracellular kinases, and identified several phosphorylation sites on AMPAR subunits (> Figure 12-1). GluR1 AMPAR subunit is phosphorylated at its C‐terminal domain in Ser831 by protein kinase C (PKC) and Ca2þ and calmodulin‐dependent protein kinase II (CaMKII) (Barria et al., 1997; Mammen et al., 1997), and in Ser845 by PKA (Roche et al., 1996). Phosphorylation of Ser845 by PKA underlies the potentiatory effect of PKA on GluR1 currents in transfected HEK 293 cells, as the effect is not observed for a Ser845 mutant (Roche et al., 1996). Further work demonstrated that PKA phosphorylation of Ser845 regulates the open channel probability of AMPAR (Banke et al., 2000). Perfusion of cells expressing GluR1 with CaMKII also resulted in a potentiatory effect on GluR1 currents, and a Ser831 to alanine mutant of GluR1 failed to be potentiated by the perfused kinase (Barria et al., 1997). Later work showed that CaMKII phosphorylation of Ser831 in GluR1 increases the single channel conductance of AMPAR (Derkach et al., 1999). Phosphospecific antibodies against the GluR1 phosphorylation sites have proven instrumental in studying GluR1 phosphorylation in vivo and in demonstrating that GluR1 phosphorylation changes during LTP and LTD (Kameyama et al., 1998; Lee et al., 1998, 2000). On the other hand, phosphorylation of GluR1 has been shown to regulate GluR1 incorporation in synapses. CaMKII drives the synaptic incorporation of GluR1‐containing AMPAR (Hayashi et al., 2000), and PKA phosphorylation of Ser845 in GluR1 is a requisite for synaptic incorporation of GluR1 mediated by CaMKII (Esteban et al., 2003). Very conclusive data were provided by studies in mice with knock in mutations in the GluR1 phosphorylation sites. The phosphomutant mice show deficits in LTD and LTP, and have memory defects in spatial learning tasks, which demonstrate that phosphorylation of GluR1 is critical for LTP and LTD expression in the hippocampus and for the retention of memories (Lee et al., 2003). GluR4 AMPAR subunit is phosphorylated at Ser842 within its C‐terminal, both in vitro and in transfected cells (Carvalho et al., 1999). PKA, PKC, and CaMKII can phosphorylate this site in GluR4. Thr830 in GluR4 is also phosphorylated by PKC in vitro (Carvalho et al., 1999). Early in the postnatal development of the hippocampus, AMPAR containing the GluR4 subunit are delivered to synapses in an activity‐dependent manner that requires PKA activity (Zhu et al., 2000). It has recently been found that activity‐driven PKA phosphorylation of GluR4 in Ser842 is necessary and sufficient to relieve a retention interaction and to drive GluR4‐containing receptors into synapses (Esteban et al., 2003), and phosphorylation of GluR4 by PKA was shown to increase the surface expression of GluR4 in cultured retina neurons (Gomes et al., 2004). Interestingly, the GluR4 phosphorylation site is conserved in GluR2long, a splice variant of GluR2 which is abundant in the hippocampus at the second postnatal week, and which delivery to the synapse is driven by spontaneous synaptic activity and during LTP (Kolleker et al., 2003). GluR4 interacts directly with PKCg through a C‐terminal membrane proximal region, and PKCg bound to GluR4 preferentially phosphorylates GluR4 in Ser842, in detriment of other substrates (Correia et al., 2003). The GluR2 subunit is phosphorylated by PKC on its C‐terminal domain, at Ser880 (Matsuda et al., 1999; Chung et al., 2000; McDonald et al., 2001), a phosphorylation site that is conserved in the other short AMPAR subunits (GluR3 and GluR4c). Moreover, Ser880 is part of the PDZ (PSD‐95, DLG, ZO‐1) domain‐ binding motif at the extreme C‐terminus of the short AMPAR subunits. PKC phosphorylates Ser880 both in vitro and in vivo, and phosphorylation of Ser880 in GluR2 plays a role in the differential binding of

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GluR2 to interacting PDZ‐domain containing proteins, glutamate receptor‐interacting protein (GRIP) and protein that interacts with C kinase (PICK1) (Matsuda et al., 1999; Chung et al., 2000), and is a critical means to regulate receptor trafficking. Interestingly, phosphorylated GluR2 can bind PICK1 but not GRIP, and is rapidly internalized (Perez et al., 2001). AMPAR endocytosis and LTD are associated with increased phosphorylation at Ser880 of GluR2 (Matsuda et al., 1999; Chung et al., 2000, 2003; Kim et al., 2001; Perez et al., 2001). GluR2 is also phosphorylated by PKC at Ser863 (McDonald et al., 2001), in vitro, and also in primary cultures of rat cortical neurons; however, the functional relevance of the phosphorylation of Ser863 has not been addressed. The GluR2 subunit is tyrosine phosphorylated on its C‐terminal domain (Ahmadian et al., 2004; Hayashi and Huganir, 2004). Ahmadian and colleagues found that tyrosine phosphorylation of GluR2 is necessary for insulin‐ and low‐frequency stimulation induced LTD of AMPAR‐mediated currents in hippocampal slices. Hayashi and Huganir identified Tyr876 in GluR2 as a phosphorylation site for the Src family of protein tyrosine kinases (PTKs), which include kinases such as Lyn, Src, and Fyn (Hayashi and Huganir, 2004). These investigators showed that phosphorylation of Tyr876 in GluR2, which is stimulated by the activation of glutamate receptors, reduces the interaction of GluR2 with GRIP, regulates AMPAR surface expression and synaptic targeting of GluR2, and is required for regulated internalization of AMPAR (Hayashi and Huganir, 2004).

2.4

AMPA Receptor Trafficking

AMPAR are crucial determinants of postsynaptic excitability, and, therefore, their density at the postsynaptic membrane is finely tuned through diverse mechanisms. AMPAR are synthesized in the endoplasmic reticulum (ER), where subunit dimerization occurs. A large pool of GluR2 remains in the ER, and RNA editing at the Arg607 (the Q/R site) in the ion pore of GluR2 controls ER retention of GluR2, ensuring the availability of GluR2 for assembly into AMPAR (Greger et al., 2002). Indeed, in hippocampal principal neurons most AMPAR are heteromeric and contain GluR2. After transiting through the Golgi, AMPAR are trafficked to dendrites or to axons. In dendrites, AMPAR exist in cytosolic vesicles or on the plasma membrane, at either synaptic or extrasynaptic sites. An unresolved issue is whether AMPAR are directly inserted into the synapse or whether their insertion is into extrasynaptic sites at the plasma membrane, from where they later diffuse to synapses. In cultured neurons, single receptor tracking techniques have shown that, although AMPAR move freely in the extrasynaptic plasma membrane, they are relatively immobile when they reach a synaptic region (Borgdorff and Choquet, 2002; Groc et al., 2004). The authors of these studies suggested that receptor diffusion between synaptic and extrasynaptic domains is involved in regulating the strength of the glutamatergic synapse, and Passafaro and colleagues (2001) found that, in fact, GluR1‐containing receptors initially accumulate at extrasynaptic sites in cultured neurons, whereas GluR2 receptors are directly inserted into the synapse. A recent study reported that AMPAR are exclusively transported from recycling endosomes to the plasma membrane during LTP of synapses (Park et al., 2004). A series of very interesting experiments established general trafficking mechanisms for AMPAR subunits (for a review see, Malinow and Malenka, 2002). By electrophysiologically tagging AMPAR subunits, to monitor synaptic delivery of recombinant AMPAR in infected neurons in hippocampal slices, Malinow and collaborators have shown that different AMPAR subunits are responsible for LTP of synapses and for receptor turnover. Receptors composed of GluR2 and GluR3 (receptors with short cytoplasmic tails) are continuously delivered to the synapse, by exchange with existing synaptic AMPAR, in an activity‐independent manner (Shi et al., 2001). In contrast, the AMPAR containing long forms of AMPAR subunits (GluR1, GluR4, and GluR2long) are restricted from accessing synapses in the absence of LTP (for GluR1‐containing receptors, Shi et al., 2001) or spontaneous activity (for GluR4‐, Zhu et al., 2000; for GluR2long‐containing receptors, Kolleker et al., 2003). After their synaptic delivery, at some point, receptors containing GluR1, GluR4, or GluR2long are replaced by GluR2/GluR3 receptors. Using a cell biology approach in dissociated cultures, Passafaro and colleagues (2001) corroborated these conclusions. On the other hand, a recent work from the laboratory of Morgan Sheng established the role of individual subunits in the removal of AMPAR from the synapses, and the subunit rules that determine the intracellular sorting of the receptors after they are

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internalized (Lee et al., 2004), and found that GluR2 is the key subunit that controls cycling and degradation of AMPAR after internalization. It is interesting to note that the distinct trafficking behaviors of short and long forms of AMPAR subunits are determined by their C‐terminal cytoplasmic tails. Proteomics studies as well as the use of the yeast two‐hybrid screening have revealed that AMPAR interact, mainly through their C‐termini, with a number of intracellular proteins that accumulate at the postsynaptic densities (> Figure 12-2; for a review see Bredt and Nicoll, 2003). These scaffolding proteins assist AMPAR traffic, or stabilize the receptors in different cellular compartments.

. Figure 12-2 Proteins that associate with AMPAR subunits. Most identified proteins associate with the intracellular C‐termini of AMPA receptor subunits. PDZ‐domain containing scaffolding proteins interact directly with AMPARs, through binding to the short forms of AMPAR (GRIP and PICK1) and to GluR1 (SAP‐97), or indirectly through transmembrane AMPAR regulatory proteins such as stargazing (PSD‐95). These scaffolding molecules recruit other proteins, such as GRASP for GRIP, AKAP for SAP‐97 and PSD‐95, and PKCa for PICK1. GluR2 also binds to NSF, and GluR1 and GluR4 bind to the cytoskeletal protein 4.1N. GluR4 associates directly with PKCg. Narp is an extracellular immediate-early early gene product, which binds to the extracellular N‐terminus of AMPAR subunits

2.4.1 Proteins that Interact with Short Forms of AMPA Receptor Subunits Proteins that contain PDZ domains play a major role in scaffolding membrane proteins, and AMPAR are no exception. The short forms of AMPAR (GluR2, GluR3, and GluR4c) terminate with the residues SVKI, which constitute a PDZ‐binding site. Two related proteins, GRIP (Dong et al., 1997) and AMPAR‐binding protein (ABP) (Srivastava et al., 1998), are PDZ‐domain containing proteins that bind the short forms of AMPAR. Several evidences suggest a role for GRIP/ABP in maintaining the synaptic accumulation of AMPAR, possibly by limiting their endocytosis (Osten et al., 2000). The C‐terminal tail of the short forms of AMPAR also binds the PDZ domain of PICK1 (Xia et al., 1999; Daw et al., 2000). Interestingly, GluR2 phosphorylation at Ser880 in the SVKI motif switches the binding preference of GluR2 from GRIP

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to PICK1 (Matsuda et al., 1999; Xia et al., 2000; Perez et al., 2001), an event which is required for LTD in the cerebellum (Chung et al., 2003). The C‐terminal segment of AMPAR short forms also binds to proteins, which do not contain PDZ domains, such as the N‐ethylmaleimide‐sensitive fusion protein (NSF), a hexameric ATPase with a role in membrane fusion (Nishimune et al., 1998; Osten et al., 1998; Song et al., 1998). NSF, which binds to a membrane proximal region of the C‐terminus of GluR2, is likely involved in the delivery of AMPAR to the plasma membrane at synapses, or in the synaptic stabilization of AMPAR, because when the interaction between NSF and GluR2 is disrupted there is rundown of excitatory postsynaptic currents in neurons in few minutes (Nishimune et al., 1998; Osten et al., 1998; Song et al., 1998). Interestingly, the NSF‐binding region in GluR2 partially overlaps the binding site for the clathrin adaptor AP2 (Lee et al., 2002), and AMPAR endocytosis occurs via a dynamin‐dependent, clathrin‐ mediated pathway, and is potentiated by factors that induce synaptic depression (Lin et al., 2000).

2.4.2 Proteins that Interact with Long Forms of AMPA Receptor Subunits GluR1 extreme C‐terminus sequence is ATGL, which conforms to the consensus motif for type I PDZ domain interactions. In fact, GluR1 was found to bind synapse‐associated protein 97 (SAP‐97) (Leonard et al., 1998), a member of the postsynaptic density‐95 (PSD‐95) protein family, through the second PDZ domain of SAP‐97 (Cai et al., 2002a). Surprisingly, SAP‐97 is not enriched in the postsynaptic densities, but rather it is associated with light membrane fractions. The role for SAP‐97 in regulating AMPAR remains controversial (Hayashi et al., 2000; Rumbaugh et al., 2003), but a recent report showed that overexpression of SAP‐97 potentiates synaptic transmission and drives GluR1-containing, AMPAR to synapses. Moreover, RNAi knockdown of endogenous SAP‐97 reduces surface expression of GluR1 and GluR2 and inhibits AMPAR‐mediated excitatory postsynaptic currents (Nakagawa et al., 2004), which suggest a major role for SAP‐97 in the maintenance of synaptic function. Interestingly, the A‐kinase anchoring protein (AKAP), which is a PKA‐ and protein phosphatase 2B (PP2B)‐scaffolding molecule, binds to SAP‐97, efficiently bringing both PKA and PP2B to GluR1 (Colledge et al., 2000). The long forms of AMPAR also participate in interactions with proteins which do not contain PDZ domains. GluR1 and GluR4 bind to the cytoskeletal protein 4.1N through the C‐terminus membrane proximal region (Shen et al., 2000; Coleman et al., 2003), and disruption of this interaction decreases the surface expression of GluR4 in heterologous cells, suggesting that anchoring of AMPAR to the cytoskeleton is important for receptor surface expression. Interestingly, the C‐terminal membrane proximal region of GluR4 interacts with the neuron‐specific isoform of PKC, PKCg (Correia et al., 2003), and the interaction facilitates GluR4 phosphorylation by PKC.

2.4.3 General AMPA Receptor Interactors: Transmembrane and Extracellular Proteins Stargazin is a transmembrane protein mutated in the stargazer mouse, which suffers from cerebellar ataxia and epilepsy (Letts et al., 1998). Stargazin interacts with AMPAR and regulates their synaptic targeting (Chen et al., 2000). Stargazin binds to the four AMPAR subunits, and its extreme C‐terminus sequence constitutes a PDZ‐domain binding site, through which stargazin associates with PSD‐95 (Chen et al., 2000). The cerebellar ataxia in the stargazer mouse is associated with the absence of functional AMPAR in cerebellar granule cells, either in the synapse or at extrasynaptic sites. Extrasynaptic AMPAR are recovered in the stargazer cerebellar granule cells when transfected with a stargazin mutant lacking the C‐terminus, and therefore unable do bind PSD‐95. On the other hand, synaptic targeting of stargazin and associated AMPAR requires transfection of granule cells with full‐length stargazin (Chen et al., 2000). This suggests that AMPAR require stargazin to reach the cell surface in cerebellar granule cells, and that, when stargazin interacts with PSD‐95, AMPAR are delivered to the synapse. In fact, overexpression of PSD‐95 in either dissociated or slice cultures causes a big increase in synaptic AMPAR (El‐Husseini et al., 2000; Schnell et al., 2002). The interaction of stargazin with PSD‐95 is regulated by reversible palmitoylation of PSD‐95 (El‐Husseini Ael et al., 2002).

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In other brain regions of the stargazer mouse there is normal AMPAR expression, which suggests that other proteins can mediate AMPAR traffic. A recent paper describes a family of transmembrane AMPAR regulatory proteins (TARPs) which interact with AMPAR through both extra‐ and intracellular determinants (Tomita et al., 2004) and rescue AMPAR in the stargazer mouse cerebellar granule cells (Tomita et al., 2003). Overexpression of stargazin does not alter the synaptic density of AMPAR, but causes a dramatic increase in their surface expression (Schnell et al., 2002), implying that stargazin is the limiting protein for surface expression of AMPAR, and showing that in hippocampal cultured neurons about 80% of AMPAR exist in intracellular stores, available for recruitment by stargazin. Interestingly, it was recently found that upon binding to glutamate, AMPAR detach from TARPs, which are stable at the plasma membrane, in contrast with AMPAR, which are internalized in a glutamate‐regulated manner (Tomita et al., 2004). The extracellular N‐terminus domain of AMPAR, at the region preceding the glutamate‐binding site (> Figure 12-1), binds to the extracellular immediate‐early gene product neuronal activity‐regulated pentraxin (Narp) (O’Brien et al., 1999), which expression is regulated by synaptic activity. Narp concentrates at excitatory synapses, particularly on the dendritic shafts of aspiny neurons (Mi et al., 2002), and Narp overexpression increases the number of excitatory synapses on dendritic shafts (O’Brien et al., 2002), and results in clusters of both NMDA‐ and AMPA‐type receptors on hippocampal interneurons. As Narp displays no ability to directly aggregate NMDA‐receptors, this effect is mediated through cytoplasmic coupling to synaptic AMPAR. It was recently found that synaptic targeting of NMDA‐receptors in aspiny spinal cord neurons, upon exogenous Narp exposure, depends on the expression of NR2A and NR2B, and requires a linkage to AMPAR through stargazin (Mi et al., 2004). Mi and colleagues suggested that in neurons which receive excitatory synapses on their dendritic shaft, such as hippocampal interneurons and spinal cord neurons, Narp plays a major role in synaptic stabilization of inserted AMPAR, whereas stargazin would play a role in bridging AMPAR to synaptic NMDA‐receptors.

3

Regulation of AMPA Receptors by Metabotropic Receptors

Metabotropic receptors are a large family of receptors for neurotransmitters and hormones that reside in the plasma membrane of neurons and glial cells, playing a modulatory role in the nervous system. These receptors are coupled to the stimulation of different G‐proteins that specifically change the activity of intracellular effector proteins. Activation of metabotropic receptors is thought to dissociate the GTP‐bound a subunits of G‐proteins from the bg subunits, and both components interact and regulate the activity of effector molecules, thereby changing the intracellular levels of cAMP, cGMP, inositol phosphates, calcium, and arachidonic acid. G‐proteins may also regulate the activity of voltage‐gated calcium channels and potassium channels (Koenig, 2004).

3.1

Metabotropic Glutamate Receptors

Based on amino acid sequence homology, pharmacological properties and the signal transduction pathways to which they are coupled, the metabotropic glutamate receptors (mGluRs) are classified into three groups: group I receptors (mGluR1 and mGluR5) are coupled to the activation of phospholipase C, whereas receptors belonging to groups II (mGluR2 and mGluR3) and III (mGluR4, mGluR6, mGluR7, and mGluR8) are coupled to the inhibition of adenylate cyclase (AC) (Conn and Pin, 1997; Schoepp et al., 1999). mGluRs have been shown to play an important role in activity‐dependent modulation of synaptic strength, which is thought to underlie learning and memory formation in the brain (Abraham and Bear, 1996; Anwyl, 1999).

3.1.1 Potentiation of AMPA Receptor by mGluRs Group I mGluRs potentiate AMPA responses in rat spinal motoneurons (Ugolini et al., 1997, 1999) and dorsal horn neurons (Bleakman et al., 1992; Cerne and Randic, 1992). mGluR1 and mGluR5 receptors are

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differentially expressed in rat motoneurons (Anneser et al., 1999; Alvarez et al., 2000) and, when stimulated with (R,S)‐3,5‐dihydroxyphenylglycine (DHPG), potentiate AMPA‐evoked membrane depolarization, through activation of PKC (Ugolini et al., 1997). However, PKC is not required for the potentiation of AMPAR activity by mGluR5 receptors, which suggests that when mGluR1 and mGluR5 are activated simultaneously the signaling cascade triggered by the former type of receptor predominates (Ugolini et al., 1999). 1S,3R‐1‐amino‐1,3‐cyclopentanedicarboxylic acid (1S,3R‐ACPD), an agonist of group I and group II mGluRs, also potentiates AMPA responses in a subpopulation of neurons from the rat visual cortex (Wang and Daw, 1996; Wang et al., 1998), in nucleus of the rat tractus solitarius (Glaum and Miller, 1993) and in rat cerebellar Purkinje neurons (Glaum etal., 1992; see > Section 3.1.2 for conflicting results), but the identity of the receptors and the signaling mechanism remain to be determined.

3.1.2 Inhibition of AMPA Receptors by mGluRs 3.1.2.1 Hippocampal Neurons Activation of group I mGluRs induces internalization of GluR1 subunits and synaptic loss of GluR1 and GluR2 in cultured hippocampal neurons (Snyder et al., 2001; Xiao et al., 2001). The internalization of AMPAR may account, at least in part, for the LTD of synaptic transmission at the Schaffer collateral‐CA1 synapses caused by group 1 mGluRs in hippocampal slices, particularly by mGluR5 (Xiao et al., 2001; Zho et al., 2002; Huang et al., 2004). A model was proposed, according to which stimulation of these receptors leads to the activation of phospholipase C through the activation of Gq proteins. The Gbg subunits released from the activated G‐protein stimulate p38 MAPK (mitogen‐activated protein kinase) through a mechanism involving Rap1, a small GTPase, and activation of MAPK kinase 3/6 (MKK3/6). Activated p38 MAPK is thought to stimulate the formation of a GDI.Rab5 (GDI‐guanyl‐ nucleotide dissociation inhibitor) complex, an important component of the molecular machinery involved in the control of endocytosis, facilitating the reduction of the plasma membrane associated AMPAR population (Huang et al., 2004). The depression of synaptic transmission requires rapamycin‐ sensitive mRNA translation, but not transcriptional activity (Snyder et al., 2001; Zho et al., 2002), which suggests that newly synthesized proteins are involved in promoting AMPAR endocytosis in response to activation of group I mGluRs. 3.1.2.2 Neurons of the Nucleus Accumbens mGluR2–5 and mGluR7 are expressed in the rat nucleus accumbens (NAc) (Testa et al., 1994; Ohishi et al., 1995, 1998), and in vivo experiments showed that injection of mGluR agonists in the NAc modulates the release of dopamine (Ohno and Watanabe, 1995; Hu et al., 1999), locomotor activity (Kim and Vezina, 1997; Swanson and Kalivas, 2000; David and Abraini, 2002), psychostimulant‐induced sensitization (Wolf, 1998), and local EEG activity in the frontal cortex (Popoli et al., 1999). Although some of these effects may occur at a presynaptic site, a recent study showed that the group III mGluR antagonist MAP4 ((S)‐2‐amino‐2‐methyl‐4‐phosphonobutanoic acid) induces a transient potentiation of the amplitude of AMPAR‐mediated currents in rat NAc neurons, induced by flash photolysis of caged glutamate (Taverna and Pennartz, 2003). However, these findings contrast with the lack of effect of the group III mGluR agonist L‐AP4 (L‐(þ)‐2‐amino‐4‐phosphonobutyric acid) on the amplitude of miniature EPSC reported by Manzoni et al. (1997). This discrepancy may be due to the activation of nonsynaptic AMPAR when photolysis of caged glutamate is used (Bernard et al., 1997). 3.1.2.3 Cerebellar Purkinje Neurons Purkinje neurons receive excitatory inputs from climbing fibers (CFs), originated in the inferior olive, and from parallel fibers (PFs), which are axons of the cerebellar granule neurons. Each Purkinje neuron receives inputs from many PFs and a single CF, and conveys the output signals originated in the cerebellar cortex. Activation, at low frequency, of PF and CF inputs to a Purkinje neuron, for several minutes, decreases PF‐Purkinje neuron synaptic drive but does not affect the efficacy of the CF‐Purkinje neuron synapse (Ito, 1989, 2002; Carvalho et al., 2000; Centonze et al., 2001; Cai et al., 2002b). This long‐term synaptic depression in the cerebellum has been suggested to contribute to certain forms of motor learning, such as adaptation of the vestibulo‐ocular reflex (Ito, 1989; Raymond et al., 1996; Boyden et al., 2004).

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Activated PFs release glutamate which stimulates AMPA receptors in the subsynaptic membrane of Purkinje cells (Baude et al., 1994; Petralia et al., 1998), together with mGluR1 receptors (Baude et al., 1993), coupled to the activation of phospholipase C (> Figure 12-3). Stimulation of these receptors leads to the production of Ins (1,4,5)P3, which releases Ca2þ from intracellular stores, and diacylglycerol, which activates PKC. Immunohistochemistry experiments showed GluR1, GluR2, GluR2/3, and mGluR1 immunoreactivity in the synaptic zone of dendritic spines at the ganglion cell‐Purkinje neuron synapses (Baude et al., 1993, 1994; Petralia et al., 1998). Initial studies performed in cultured chick Purkinje neurons and in slices from the rat cerebellum showed that mGluRs inhibit AMPA‐mediated responses in Purkinje cells (Ito and Karachot, 1990; Mori‐Okamoto et al., 1993). Subsequent studies showed that the induction of LTD at the PFs‐ Purkinje cell synapses requires activation of mGluR1 (Kano and Kato, 1987; Linden et al., 1991; Aiba et al., 1994; Conquet et al., 1994; Hartell, 1994; Shigemoto et al., 1994), Ca2þ influx through voltage‐gated Ca2þ channels (Linden et al., 1991; Konnerth et al., 1992) and AMPA receptor activation (Linden et al., 1993). These three signals appear to cooperate to promote the activation of PKC (Linden and Connor, 1991), which phosphorylates GluR2 at Ser880 (Chung et al., 2003). The importance of GluR2 phosphorylation at Ser880 to cerebellar LTD was shown in studies using cultured cerebellar Purkinje cells from mutant mice lacking GluR2. These cells do not show LTD, which, however, is restored upon transfection with the wild‐ type GluR2. In contrast, transfection with a point mutant that eliminates PKC phosphorylation on Ser880 does not restore LTD (Chung et al., 2003). Phosphorylation of GluR2 on Ser880 is thought to decrease the number of synaptic AMPA receptors upon induction of LTD (Matsuda et al., 2000), via clathrin‐mediated endocytosis (Lin et al., 2000), followed by internalization of the receptors. The declustering of the receptors . Figure 12-3 Molecular mechanisms involved in the regulation of AMPAR during LTD at the parallel fiber (PF)‐Purkinje cell (PC) synapses in the cerebellum. LTD in these synapses is generated by synchronous activation of the PF and climbing fiber (CF) inputs to a Purkinje neuron

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may arise from the phosphorylation of GluR2, which disrupts GluR2 binding to GRIP1 and 2, and promotes binding to PICK1 (Matsuda et al., 1999; Chung et al., 2000; Perez et al., 2001). In addition to the phosphorylation of GluR2, PKC may also phosphorylate CPI‐17 in response to the induction of LTD in cultured Purkinje neurons (Eto et al., 2002). CPI‐17 is activated upon phosphorylation by PKC, thereby inhibiting myosin/moesin phosphatase (MMP). The resulting inhibition of the phosphatase, together with the coincident PKC activation, might increase phosphorylation of Ser880 and thereby promote the internalization of GluR2‐containing AMPA receptors. The a isoform of PKC is involved in the induction of cerebellar LTD, as shown in studies with PKCa null mice or with Purkinje cells in which the enzyme protein levels were reduced by targeted RNA interference (Leitges et al., 2004). This role of PKCa may derive from its unique PDZ‐binding motif, located at the very C‐terminal end of the protein, which is not present in the other isoforms (Leitges et al., 2004). Biochemical evidences also suggest that only active PKCa can bind PICK1 (Perez et al., 2001), suggesting that the PICK1 bound kinase may phosphorylate GluR2 at Ser 880. Although the major role of mGluR1 activation in LTD in the cerebellum appears to arise from the activation of PKC (see earlier), a role for Ca2þ release from Ins(1,4,5)P3‐sensitive stores has also been suggested (reviewed in Bear and Linden, 2001; Centonze et al., 2001). In fact, Purkinje cells express unusually high levels of Ins(1,4,5)P3 receptors, particularly the type I isoform (Nakanishi et al., 1991), but not all experimental evidences support their role in LTD (Bear and Linden, 2001; Centonze et al., 2001). The glutamate receptor d2 (GluRd2) is selectively expressed in Purkinje cells, where it is found at the postsynaptic side of the PFs‐Purkinje cell synapses (Araki et al., 1993; Lomeli et al., 1993; Landsend et al., 1997). Although GluRd2 does not form functional glutamate‐gated ion channels, mutant mice deficient in the protein show impairments in LTD at the PFs‐Purkinje neuron synapses (Kashiwabuchi et al., 1995) and in motor learning (Funabiki etal., 1995; Kishimoto etal., 2001, Yuzaki, 2004). A recent study showed that GluRd2 interacts with Shank proteins through an internal motif. Immunoprecipitation studies with an antibody against GluRd2 also revealed an interaction with Homer and mGluR1 in synaptosomal membrane fractions (Uemura et al., 2004). Since Shank2 also interacts with GRIP1 in the cerebellum, it was suggested that under resting conditions GluRd2 participates in a complex with mGluR1 and AMPA receptors at the synapses between PFs and the dendrites of Purkinje neurons (Uemura et al., 2004). However, it remains to be determined how GluRd2 contributes to the development of LTD in these synapses. Although genetic ablation of the neuronal isoform of nitric oxide synthase (nNOS) impairs cerebellar LTD (Lev‐Ram et al., 1997) and adaptation of compensatory eye movements (Katoh et al., 2000), the molecular mechanism whereby nitric oxide (NO) affects LTD in the cerebellum is still not completely understood. The nNOS is not found in Purkinje cells in vivo (Crepel et al., 1994), suggesting that the effect of NO may be due to the activity of the enzyme present in neighboring PFs (Shibuki and Kimura, 1997), in response to a rise in the [Ca2þ]i. NO is thought to activate guanylate cyclase in Purkinje neurons (Ariano et al., 1982; Boxall and Garthwaite, 1996), giving rise to cGMP which stimulates cGMP‐dependent protein kinase (PKG) (Ito and Karachot, 1992). Cerebellar Purkinje neurons contain large amounts of PKG type I (Hofmann and Sold, 1972; Lohmann et al., 1981) and conditional knockout mice lacking PKG type I selectively in Purkinje neurons show a strong reduction in LTD and an impaired adaptation of the vestibular‐ocular reflex (Feil et al., 2003). However, it remains to be determined how PKG affects AMPA receptor mediated responses. One of the substrates of this kinase is substrate G, which is expressed in the cerebellum and inhibits protein phosphatase 1/2A activity in its phosphorylated form (Detre et al., 1984; Endo et al., 1999; Hall et al., 1999). Accordingly, inhibition of protein phosphatase 2A induces cerebellar LTD in cultured Purkinje cells and causes the declustering of synaptic AMPA receptors (Launey et al., 2004). Protein phosphatase 2A may directly dephosphorylate GluR2, as recently suggested based on in vitro studies, or may act as a regulator of the activity of PKC (Launey et al., 2004).

3.2

Dopamine Receptors

Dopamine modulates glutamate transmission within multiple brain regions, playing a role in the normal regulation of motor, affective, and cognitive functions. The majority of dopaminergic neurons are localized to the substantia nigra and to more diffuse cell groups in the ventral tegmental area (VTA). Smaller

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groups of dopaminergic neurons are found in the retina, in the hypothalamus, and in the olfactory bulb. Dopamine is a monoamine, which acts through a group of metabotropic receptors divided into two families based on pharmacological and biochemical criteria (Missale et al., 1998). Receptors belonging to the D1‐like family (D1 and D5 receptors) are coupled to Gs/olf proteins and activate AC, whereas the receptors that constitute the D2‐like family (D2–D4) activate Gi/o proteins, and therefore inhibit AC. There is also evidence suggesting that the D1‐like receptors may stimulate the phosphoinositide turnover pathway (Mahan et al., 1990; Rodrigues Pdos and Dowling, 1990; Undie and Friedman, 1990, 1992; Undie et al., 1994; Friedman et al., 1997). The most prominent dopaminergic pathway in the central nervous system is the nigrostriatal pathway; dopamine modulates the response of striatal neurons to excitatory inputs from the cerebral cortex and the thalamus. Ionotropic glutamate receptors mediate synaptic transmission and plasticity in the striatum, whereas dopamine receptors present both pre‐ and postsynaptically modulate the excitability of the medium spiny neurons by coupling to G‐protein pathways. Scattered dopaminergic cells in the VTA dorsal to the substantia nigra send dopaminergic afferents to the ventral striatum and to the frontal lobes. Imbalances in dopaminergic and glutamatergic synaptic transmission in the striatum, NAc, and prefrontal cortex have been implicated in several disorders, including Parkinson’s disease, Huntington’s disease, schizophrenia, and drug addiction. Three levels of interaction between dopamine and glutamate exist. Dopaminergic neurons can innervate glutamatergic neurons and regulate glutamate release, whereas glutamatergic neurons can modulate dopamine release from dopaminergic neurons. In this chapter, we focus on a third level of interaction between the two systems, concerning the modulation of AMPAR by dopamine receptors, via convergent connections onto common dendritic targets.

3.2.1 Striatum A complex interaction between glutamatergic and dopaminergic systems in the dorsal striatum and NAc is involved in the gating of information flow in these integrative brain regions. In the striatum, morphological evidence for the convergence of glutamatergic and dopaminergic inputs is well established for cortical afferents. It was first identified in the dorsal striatum, where dopaminergic terminals form synapses in spines from medium spiny GABAergic projection neurons, which receive glutamate synaptic input from the sensorimotor cortex (Bouyer et al., 1984; Freund et al., 1984; Smith et al., 1994; Hidaka and Totterdell, 2001). This convergent arrangement has now been demonstrated for the ventral striatum: the NAc receives cortical inputs from the hippocampus and the amygdala, which converge with dopaminergic terminals on spiny neurons and on the aspiny dendrites of interneurons (Totterdell and Smith, 1989; Sesack and Pickel, 1990; Johnson et al., 1994; Hidaka and Totterdell, 2001). The thalamic nuclei are another important source of glutamate inputs to the striatum. An early report showed no convergent synaptic relationship between thalamus afferents and dopaminergic terminals (Smith et al., 1994), but more recent reports demonstrated postsynaptic convergence between dopamine and thalamic afferents to a striatal region (LeDoux et al., 1990; Campeau and Davis, 1995; Pinto et al., 2003). Ultrastructural studies have shown that dopaminergic terminals appose both symmetrical and asymmetrical synapses (Sesack and Pickel, 1992), and a portion of dopamine inputs do not make classical synaptic contacts onto postsynaptic structures (Bouyer et al., 1984; Descarries et al., 1996). Thus, dopaminergic afferents to the dorsal striatum and to the NAc modulate synaptic transmission by releasing dopamine in the vicinity of glutamatergic synapses or by causing diffuse increases in extracellular dopamine levels (Garris et al., 1994; Descarries et al., 1996; Gonon, 1997). Moreover, glutamatergic and dopaminergic transmission can interact via both pre‐ and postsynaptic mechanisms. In striatal spiny neurons, dopamine and glutamate receptors colocalize (Ariano et al., 1997), and, on the other hand, glutamate release is modulated by dopamine (Brown and Arbuthnott, 1983), whereas dopamine release is modulated by glutamate (Moghaddam et al., 1990). In this chapter, we focus on the interactions between glutamate and dopamine systems at the postsynaptic level, whereas others have summarized the literature on the interactions between the two systems at the presynaptic level (Morari et al., 1998).

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3.2.1.1 Dorsal Striatum 3.2.1.1.1 Modulation of Synaptic Transmission by Dopamine in the Dorsal Striatum Excitatory cortical

afferents evoke in striatum medium spiny neurons a dual component postsynaptic potential, mediated by AMPA‐ and NMDA‐receptors (Uchimura et al., 1989; Jiang and North, 1991; Kombian and Malenka, 1994). Several evidences suggest that, in the striatum, dopamine can have distinct effects on the two types of receptors depending on the subtype of dopamine receptor that is activated. We focus on the effects of dopamine on AMPAR‐mediated postsynaptic potentials. Several studies reported D1‐receptor mediated enhancement of AMPAR excitatory postsynaptic potentials (Umemiya and Raymond, 1997; Lin et al., 2003), and prevention of the rundown of iontophoretic AMPAR currents, which occurs in dissociated medium spiny cells (Yan et al., 1999). The mechanism for this latter effect has been shown to involve activation of PKA and phosphorylation of DARPP‐32 (dopamine‐ and cAMP‐regulated phosphoprotein, Mr 32,000) (Yan et al., 1999), a dopamine‐ and cAMP‐regulated phosphoprotein highly enriched in all dopamine‐innervated neurons (Walaas et al., 1983). Yan and colleagues showed that, in dissociated medium spiny neurons, D1 receptor activation, protein phosphatase 1/2A inhibition, intracellular perfusion with a phospo‐DARPP‐32 peptide, or disruption of the interaction between protein phosphatase‐1 (PP‐1) and spinophilin can stabilize AMPA currents. The authors of this study suggested that PP‐1 is anchored to AMPAR at the postsynaptic density through spinophilin, which facilitates the ability of PP‐1 to dephosphorylate AMPAR. When D1 receptors are activated, PKA activity is increased and DARPP‐32 is phosphorylated. Phosphorylated DARPP‐32 inhibits PP‐1, which dissociates from spinophilin and is therefore removed from the vicinity of AMPAR. This cascade of events converts AMPAR to the phosphorylated active state (> Figure 12-4), through a synergistic action involving decreased dephosphorylation of AMPAR by PP‐1 (Yan et al., 1999) and direct phosphorylation of GluR1 AMPAR at Ser845 by PKA (Price et al., 1999; Snyder et al., 2000). Interestingly, the phosphorylation of the PKA phosphorylation site in GluR1 (Ser845) is strongly increased in neostriatum in vivo in response to the psychostimulants cocaine and methamphetamine, and the effects on GluR1 phosphorylation are attenuated in DARPP‐32 knockout mice, suggesting that DARPP‐32 and AMPAR are cellular effectors for psychostimulant actions (Snyder et al., 2000). A recent study showed that D1‐receptor activation leads to phosphorylation at the Ser845 site in GluR1, and to enhanced surface expression of GluR1 in cultured striatal neurons (Swayze et al., 2004). Moreover, overexpression of mutant forms of the AKAP79/150 and PSD‐95 significantly attenuates the effects of the D1‐receptor agonist on GluR1 phosphorylation and surface expression, suggesting that PSD‐95 and the associated AKAP79/150 are critical for mediating cross‐talk between the dopaminergic and glutamatergic systems (Swayze et al., 2004). . Figure 12-4 D1 dopamine receptor activation enhances AMPAR‐mediated responses in striatum medium spiny neurons. The mechanism for this effect involves activation of PKA, and direct phosphorylation of GluR1 by PKA. Moreover, PKA phophorylates DARPP‐32, which inhibits PP‐1 and leads to PP‐1 dissociation from spinophilin and its removal from the vicinity of AMPAR

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The understanding of the synaptic actions of dopamine in the striatum is complicated by the fact that several studies were unable to detect effects of dopamine on excitatory synaptic responses in striatal slices. Accordingly, D1‐receptor activation causes no alterations in AMPAR excitatory postsynaptic potentials (Levine et al., 1996), and no effects of dopamine were found on excitatory synaptic responses in field potential or whole‐cell recordings from striatal slices (Malenka and Kocsis, 1988; Nicola and Malenka, 1998). D2‐receptor activation was reported to lead to reduced AMPAR‐mediated postsynaptic potentials (Cepeda et al., 1993; Levine et al., 1996), but others found that the D2‐receptor mediated reduction of the excitatory postsynaptic potential is observed only in slices from animals subjected to 6‐hydroxydopamine lesions of the substantia nigra (Calabresi et al., 1993). Malenka’s group suggested that the explanation for these conflicting results may be that dopamine has no direct effect on excitatory synaptic transmission in the striatum (i.e., no effect on postsynaptic glutamate receptors), but rather that dopamine indirectly modulates excitatory responses via action on the voltage‐dependent calcium channels (Nicola et al., 2000). 3.2.1.1.2 Dopaminergic Control of Synaptic Plasticity in the Dorsal Striatum LTP in the striatum is observed in vivo following repetitive stimulation of ipsilateral cortical afferents (Charpier and Deniau, 1997), and is dependent on the activation of NMDA‐receptors (Calabresi et al., 1992a; Partridge et al., 2000). High‐ frequency stimulation of the contralateral cerebral cortex induces LTD, which is associated with dopamine depletion in the striatum (Reynolds and Wickens, 2000). Repetitive stimulation of corticostriatal fibers produces a dramatic increase in the release of glutamate and dopamine in the striatum, suggesting that these neurotransmitters operate together to induce striatal synaptic plasticity (for a review see, Centonze et al., 2001). Moreover, LTP and LTD disappear in the striatum when the catecholamine‐specific neurotoxin 6‐hydroxydopamine is injected into the substantia nigra of rats, causing a permanent dopamine‐denervation of the ipsilateral striatum (Calabresi et al., 1992b; Centonze et al., 1999). In vitro studies have found that stimulation of dopamine receptors is a critical requirement for corticostriatal LTD. Both antagonists of the D1‐ and D2‐like receptors prevent striatal LTD, which is absent in the striatum of mice lacking D2‐receptors (Calabresi et al., 1992b, 1997). D1‐ and D2‐receptors cooperate to induce LTD in the striatum, through a mechanism involving a presynaptic effect and NO production by the striatal interneurons (Centonze et al., 2001). In contrast, LTP in the striatum is enhanced by D2‐receptor antagonists and in mice lacking D2‐receptors, and the activation of D2‐receptors fully prevents LTP (Calabresi et al., 1997). On the other hand, D1‐receptor or postsynaptic PKA inhibition results in blockade of LTP in the striatum (Calabresi et al., 2000; Kerr and Wickens, 2001). Together, these evidences indicate that PKA stimulation, via postsynaptic D1‐like receptor activation or D2‐like receptor inhibition, favors corticostriatal LTP. A major substrate for PKA in the spiny neurons is DARPP‐32, and in DARPP‐32 knockout mice both corticostriatal LTD and LTP are absent (Calabresi et al., 2000). Pharmacological inhibition of PP‐1 is able to restore both forms of plasticity, indicating that the inhibition of PP‐1 by phosphorylated DARPP‐32 is crucial for the induction of LTP and LTD in the striatum. LTP induction is dependent on the phosphorylation of DARPP‐32 by postsynaptic PKA, activated following D1‐receptor activation. The inhibition of PP‐1 by phosphorylated DARPP‐32 results in decreased dephosphorylation of many physiological effectors, including NMDA and AMPAR, which are PP‐1 substrates (Greengard et al., 1999). 3.2.1.2 Nucleus Accumbens The NAc is an interface between the limbic system, which controls motivational aspects of behavior, and the motor systems, which control the execution of behavior. The neurons in this brain region receive convergent inputs from the VTA‐dopamine containing neurons and from glutamate‐containing neurons originating from cortical and limbic brain regions, such as the prefrontal cortex, the hippocampus, and the amygdala (Sesack and Pickel, 1990; Groenewegen et al., 1999). The most abundant population of neurons in the NAc is constituted by medium spiny GABAergic neurons, which are the projection neurons in this brain region. The remaining
10% of cells is composed of several populations of interneurons. The NAc is a critical brain region for drug addiction and it is the brain region where many drugs induce alterations in glutamate transmission (for a review see, Wolf, 1998). In fact, in vivo exposure to drugs of abuse such as amphetamine and cocaine, which target the dopamine

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transporter and lead to an acute increase in synaptic dopamine levels, has been shown to elicit changes in synaptic strength at excitatory synapses, producing LTD‐like effects in the NAc (Thomas et al., 2001) and LTP‐like effects in the VTA (Ungless et al., 2001; Saal et al., 2003). The cocaine‐induced synaptic enhancement in the VTA involves an upregulation of AMPAR, which requires GluR1, as it is absent in mice lacking this subunit (Dong et al., 2004). Dopamine applied iontophoretically has a marked attenuating effect on the excitatory response of NAc neurons to amygdala stimulation (Yim and Mogenson, 1982) or to hippocampal stimulation (Yang and Mogenson, 1984), at low frequency (DeFrance et al., 1985). These early results, obtained in vivo, suggested that dopamine acts in the NAc to increase the ‘‘signal‐to‐noise’’ ratio, which might be a form of ‘‘contrast enhancement’’ of an incoming hippocampal message (DeFrance et al., 1985). Endogenous release of dopamine caused by stimulation of the VTA also attenuated the response of NAc neurons to excitatory inputs from the amygdala (Yim and Mogenson, 1982) or the hippocampus (Yang and Mogenson, 1984). Moreover, a recent in vivo study showed that the EPSP elicited in the NAc neurons by prefrontal cortex stimulation is attenuated by stimulation of the VTA, through a D2‐receptor mediated effect (Brady and O’Donnell, 2004). However, another study reported that either chemical stimulation of the VTA, using NMDA microinjection, or burst firing of dopamine neurons, enhances the response of NAc neurons. This excitation is sensitive to antagonists of the D1 dopamine receptors. Moreover, excitatory responses in the NAc to electrical stimulation of the hippocampus are strongly facilitated by endogenously released dopamine (Gonon and Sundstrom, 1996). The effects of dopamine in the NAc neurons are complex, and both excitatory and inhibitory effects of dopamine on the synaptic activity of medium spiny neurons, evoked by excitatory synaptic afferents, have been reported (see earlier). Therefore, recent studies tested the possibility that these opposing actions of dopamine play a role in the integration and gating of different limbic signals to the NAc. For that purpose, Floresco and colleagues (2001b) have performed electrophysiological recordings in vivo from NAc neurons that responded to hippocampal stimulation alone or to both hippocampal and amygdala stimulation. Their data suggest that dopamine transmission in the NAc plays a critical role in an input selection mechanism, permitting certain inputs to have preferential temporary influence over neural activity in the NAc. They found that high‐frequency activity in the glutamatergic hippocampal‐NAc pathway can facilitate the release of dopamine, which facilitates subsequent hippocampal‐driven neural activity in NAc neurons, through a D1‐receptor mediated effect. On the other hand, the dopamine release triggered by high‐frequency hippocampal inputs has an inhibitory effect on the amygdala inputs on the same NAc neuron (Floresco et al., 2001b). A further study showed that, when dopamine release happens following high‐frequency stimulation of amygdala inputs to the NAc, this is correlated with an enhancement of firing of NAc neurons in response to amygdala stimulation, an effect that is mediated by D1‐receptors and NMDA‐receptors (Floresco et al., 2001a). Taken together, these results indicate that the modulation of dopamine release may serve as an important component of a gating mechanism of ensuring that a specific input to the NAc has preferential influence on neuronal activity (Floresco et al., 2001a, b), and may help reconcile apparently contradictory previous reports. In vitro studies using slice preparations investigated the dopamine effects on excitatory synaptic transmission in the NAc and found a D1‐receptor mediated depression of excitatory synaptic responses, which appears to be due to a presynaptic effect (Pennartz et al., 1992; Harvey and Lacey, 1996, 1997; Nicola et al., 1996; Nicola and Malenka, 1997, 1998). The role of the NAc in drug addition has been the subject of intensive investigation in the recent years. The cellular mechanisms that underlie the postsynaptic interactions between dopamine and glutamate receptors on medium spiny neurons, which are probably critical for acute and chronic effects of cocaine and amphetamine, were addressed in NAc cell cultures (Chao et al., 2002a, b; Mangiavacchi and Wolf, 2004). In this preparation, D1 receptor stimulation increases the phosphorylation of the AMPAR subunit GluR1 at the PKA phosphorylation site (Ser845), whereas stimulation of D2 receptors attenuates the response to D1 receptor activation (Chao et al., 2002b). Phosphorylation of Ser845 in GluR1 by PKA increases AMPAR

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peak response open probability (Banke et al., 2000), and is critical for synaptic incorporation of GluR1‐ containing AMPAR and for LTP in the hippocampus (Esteban et al., 2003). In fact, D1 receptor stimulation increases GluR1 surface expression in cultured NAc neurons (Chao et al., 2002a), by increasing the rate of AMPAR insertion onto the surface of cultured NAc neurons (Mangiavacchi and Wolf, 2004). Because the NAc cultures do not contain glutamatergic neurons, GluR1 is incorporated, following D1 receptor stimulation, into extrasynaptic sites. This is suggested to constitute a possible mechanism by which amphetamine and cocaine, which indirectly stimulate D1 receptors, alter synaptic strength in addiction‐ related neuronal circuits. However, in NAc slices prepared from cocaine‐treated animals, excitatory synapses showed a decrease in synaptic strength, and diminished LTD, which indicates that the decrease in synaptic strength is due to mechanisms shared with LTD (Thomas et al., 2001).

3.2.2 Prefrontal Cortex The prefrontal cortex is an important brain region in which to study dopamine and glutamate receptor interactions, because these interactions were shown to play a role in disorders involving the dysfunction of the prefrontal cortex, such as schizophrenia (Laruelle et al., 2003) and drug addiction (Dackis and O’Brien, 2003). The prefrontal cortex receives inputs from cortical and subcortical areas involved in sensorimotor and limbic functions, and from dopamine‐containing neurons in the VTA, which projects to the prefrontal cortex. A recent study using brain slices from mature rats examined the cellular mechanism involved in the modulation of AMPA responses by D1 and D2 receptors in prefrontal cortex pyramidal neurons, and found that D2 receptors inhibit AMPA responses, through two different postsynaptic pathways, one that requires intracellular Ca2þ and the phospholipase C‐Ins(1,4,5)P3 cascade, the other requiring the blockade of the cAMP‐PKA cascade (Tseng and O’Donnell, 2004). This study found no interaction between D1 receptors and AMPAR (Tseng and O’Donnell, 2004), but an in vitro study in layer II–III pyramidal cells showed that bath application of dopamine significantly enhances AMPA‐mediated EPSC amplitudes (Gonzalez‐Islas and Hablitz, 2003). Application of a specific D1‐like receptor agonist significantly increased EPSC amplitude, whereas the D2‐like receptor agonist quinpirole had no effect (Gonzalez‐Islas and Hablitz, 2003). In primary postnatal prefrontal cortex cultures, activation of D1 receptors increases surface expression of GluR1 (Wolf et al., 2003; Swayze et al., 2004), but not synaptic insertion of GluR1‐containing AMPAR (Wolf et al., 2003). The authors suggested that D1 receptor stimulation increases the nonsynaptic cell surface pool of GluR1 in prefrontal cortex neurons, and thus increases the GluR1 pool available for insertion in the synapse during LTP. Accordingly, D1 receptors have been shown to promote LTP in the prefrontal cortex (Gurden et al., 2000).

3.2.3 Retina In the retina, dopamine is released by a set of dopaminergic amacrine/interplexiform neurons. Retinal dopamine levels are rhythmic, and regulated by a combination of circadian rhythmicity and light exposure; dopamine release is lower in darkness and higher in light, the time when vision switches from being rod‐mediated to being cone‐mediated, and therefore becomes less sensitive but more acute (Witkovsky, 2004). Bipolar, horizontal, amacrine and ganglion cells have D1 receptors (Veruki and Wassle, 1996; Nguyen‐Legros et al., 1997), whereas the retinal pigment epithelium has D5 receptors (Versaux‐Botteri et al., 1997). Rod and cone photoreceptors (Muresan and Besharse, 1993), as well as dopaminergic cells (Veruki, 1997), express D2 family receptors. The effect of dopamine in the outer retina was initially studied in vitro using fish horizontal cells in culture (Knapp and Dowling, 1987). Dopamine application increases the response to kainate, by increasing the probability of channel opening for a given agonist concentration without changing the amount of current passed by an individual channel (Knapp et al., 1990). The mechanism is dependent on D1 receptors

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and on the cAMP/PKA cascade. Interestingly, phosphorylation of Ser845 in GluR1 by PKA was later found to increase AMPAR peak response open probability (see earlier, Banke et al., 2000). However, another study in retinal horizontal cells of the perch suggested that the dopamine‐dependent modulation of glutamate receptors is due to a reduction in the desensitization of glutamate receptors (Schmidt et al., 1994). The enhancing action of dopamine appeared to be specific to cone horizontal cells, because kainate‐induced currents of rod horizontal cells, which lack dopaminergic innervation in vivo, failed to increase following treatment with dopamine (Knapp and Dowling, 1987). In fact, an in vivo study showed that dopamine increases the cone input and suppresses the rod input to horizontal cells of the Xenopus retina, through an effect dependent on D1 receptors (Witkovsky et al., 1988). Another study found that dopamine may modulate synaptic transmission from photoreceptors to OFF bipolar cells (Maguire and Werblin, 1994). Bath‐applied dopamine enhanced the response to glutamate in OFF bipolar cells in retinal slices and in isolated OFF bipolar cells. The enhancement by dopamine is mediated by D1 receptors, and blocked by a PKA inhibitor. These data suggest that dopamine acts at D1 receptors in the dendrites of bipolar cells to activate AC, which through cAMP enhances a glutamate‐gated current. In the inner retina, amacrine cells receive glutamatergic input from bipolar cells and make synapses on the ganglion cells as well as inhibitory synapses on the axon terminal of bipolar cells, thus controlling their output to ganglion cells. Amacrine cells have dedicated functions recently reviewed extensively (Masland, 2001). Dopaminergic amacrine cells adjust the retina’s responsiveness under bright or dim light (see earlier), whereas starburst amacrine cells make excitatory cholinergic synapses on ganglion cells sensitive to moving stimuli, playing an important role in direction selectivity; polyaxonal amacrine cells play a role in distinguishing motion of an object from motion across the retina caused by the fixational eye movements (Olveczky et al., 2003). An in vitro study investigated whether the amacrine cell response to kainate is modulated by dopamine. In primary cultures enriched in chick cholinergic amacrine cells, it was found that activation of D1 receptors increases the [Ca2þ]i response to kainate (Gomes et al., 2004), which is mediated by AMPAR (Carvalho et al., 1998). In these cultures, AMPAR are enriched in GluR4 (Carvalho et al., 2002), which is a substrate for PKA (Carvalho et al., 1999). Gomes and colleagues (2004) found that D1 receptor stimulation in this population of amacrine cells in culture leads to increased phosphorylation of Ser842 in GluR4, and increased GluR4 recruitment to the plasma membrane. Interestingly, the same study found that group II mGluRs cross‐talk with D1 receptors in this system, because activation of group II mGluRs, negatively coupled to AC, prevents the effect of D1 receptor activation on AMPAR activity, GluR4 phosphorylation and GluR4 surface expression (Gomes et al., 2004).

3.2.4 Regulation of AMPA Receptor Subunit Expression by Dopamine Several studies demonstrate that the glutamatergic system undergoes regulatory changes at the level of glutamate receptor expression after dopamine depletion. Lesion of the substantia nigra pars compacta, by administration of dopamine, downregulates striatal expression of GluR1 AMPAR subunit (Fan et al., 1999) and hippocampal expression of both GluR1 and GluR2 subunits (Fan et al., 2000), as determined by in situ hybridization. Other studies investigated the expression of AMPAR GluR1 subunit in the neostriatum of the 6‐hydroxydopamine‐lesioned rat, which is an animal model of Parkinson’s disease. RT‐PCR studies showed that GluR1 mRNA is reduced 2 weeks after the lesion, and immunofluorescence studies demonstrated a significant reduction in the GluR1 immunoreactivity (Lai et al., 2003). Neither the mRNA nor the protein levels for GluR2/3 or GluR4 subunits were altered. In the mesencephalon, 1 day after the 6‐hydroxydopamine lesion GluR1, GluR2, and GluR3 mRNA levels were decreased (Bardo et al., 2001). However, in D1A‐deficient mice immunostaining for GluR1 was increased in comparison with littermate controls in striatal interneurons (Ariano et al., 1998). Following nigrostriatal dopaminergic denervation, the glutamatergic corticostriatal pathway is known to become overactive. A recent study analyzed the regulation of the GluR1 subunit in the basal ganglia of primates following dopamine denervation, and found that GluR1 protein expression is increased in caudate

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and putamen areas of the striatum, altered minimally in the subthalamic nucleus, and downregulated in the globus pallidus and in the substantia nigra (Betarbet et al., 2000). 3.2.4.1 Effect of Antipsychotic Drugs on AMPA Receptor Subunit Expression The dopamine–glutamate interactions in limbic cortex and striatum are considered crucial for symptom production in schizophrenia, and AMPAR subunit expression has been found to be decreased in the medial temporal lobe in schizophrenia (Meador‐Woodruff and Healy, 2000). Moreover, antipsychotics, the mainstay of treatment of this illness, generally antagonize dopamine receptors. Therefore, various research groups have investigated whether antipsychotic drugs affect the expression of glutamate receptor subunits in the critical brain regions for schizophrenia. Antipsychotic drugs do alter the expression of the genes encoding the subunits that express ionotropic glutamate receptors. An early study found that in the striatum haloperidol has no effect on GluR1 or GluR2 protein levels, but that both haloperidol and clozapine increase GluR1 levels in the medial prefrontal cortex (Fitzgerald et al., 1995). However, haloperidol was found to cause a decrease in GluR2 and GluR4 mRNA levels in both cortex and striatum, whereas clozapine treatment causes GluR3 decrease in cortex and striatum, but a GluR4 increase in the striatum only (Healy and Meador‐Woodruff, 1997). Another study found that haloperidol causes a small but significant induction of GluR2 flip mRNA in the dorsolateral caudate putamen (Brene et al., 1998). 3.2.4.2 Effect of Drugs of Abuse on AMPA Receptor Subunit Expression There is compelling evidence that the VTA dopamine system and its glutamate‐mediated inputs are crucial for the development of sensitized responses to drugs of abuse. The NAc, innervated by the VTA, appears to mediate the expression of sensitization (Nestler, 2004). Repeated intermittent administration of drugs of abuse, such as cocaine and amphetamines, which are indirect agonists at dopamine receptors by inhibiting dopamine transporters or stimulating dopamine release, respectively, causes numerous alterations in the levels of different proteins within the VTA, including changes in AMPAR subunits. Several studies found that GluR1 protein expression in the VTA is increased by repetitive cocaine treatment (Fitzgerald et al., 1996; Churchill et al., 1999). The authors of these studies suggested that increased GluR1 expression in the VTA may represent a molecular mechanism by which drugs of abuse exert long‐term effects on dopaminergic function. However, another study could not find changes in GluR1 at the protein or at the mRNA level in the VAT, after cocaine or amphetamine administration (Lu et al., 2002). Nonetheless, the effects of drugs of abuse on GluR1 expression could be of interest, because elevated levels of GluR1 would favor the formation of Ca2þ‐permeable AMPAR, which would trigger Ca2þ‐dependent signaling cascades within the VTA (Carlezon and Nestler, 2002). In this review, Carlezon and Nestler propose that drug sensitization involves pathophysiological mechanisms that can be triggered by processes involving accumulation of GluR1 in the VTA. GluR1‐homomeric, Ca2þ‐permeable, AMPAR would lead to increases in intracellular [Ca2þ], which would cause widespread alterations in gene expression, accompanied by transient increases in the excitability and function of VTA dopaminergic neurons. These changes would augment the sensitivity of these neurons to the pharmacological actions of drugs of abuse (Carlezon and Nestler, 2002). However, a recent study showed that, surprisingly, behavioral sensitization to cocaine can be elicited in GluR1(/) mice (Dong et al., 2004). The data obtained by Dong and colleagues further show that GluR1(/) mice do not exhibit conditioned place preference in response to cocaine, suggesting that the drug‐induced enhancement of excitatory synaptic transmission in DA neurons, which is dependent on GluR1 expression, although not required for drug sensitization may contribute to attribution of incentive value to drug‐associated references. Drug treatment was shown to alter AMPAR subunit expression in other brain regions. In the NAc, acute cocaine treatment significantly reduces the mRNA level for GluR3 and GluR4 (Ghasemzadeh et al., 1999), and repeated amphetamine administration decreases levels of GluR1 and GluR2, but not GluR3 mRNAs (Lu et al., 1997). In the prefrontal cortex, repeated cocaine administration increases the level of GluR2 mRNA (Ghasemzadeh et al., 1999), whereas GluR1 mRNA levels were found to be increased in this brain region after amphetamine withdrawal (Lu et al., 1997). Because functional properties of AMPAR are determined by subunit composition, these changes may result in altered AMPA transmission in NAc and

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prefrontal cortex. However, it is still unclear whether these changes in AMPAR subunit composition are direct consequences of drug treatment of secondary to other changes caused by drug treatment.

3.3

Serotonin Receptors

Serotonin interacts with a large family of receptors, which have been divided into seven subfamilies based on their pharmacological properties, amino acid sequence, gene organization, and signaling mechanisms. The 5‐HT3 receptors are ionotropic receptors, and the 5‐HT1 (5‐HT1A, 5‐HT1B, 5‐HT1D‐5‐HT1F), 5‐HT2 (5‐HT2A‐5‐HT2C), 5‐HT4, 5‐HT5 (5‐HT5A, 5‐HT5B), 5‐HT6, and 5‐HT7 receptors belong to the large family of G‐Protein Coupled Receptors (GPCR) (Barnes and Sharp, 1999; Raymond et al., 2001). Although there are exceptions to the general pattern of G‐protein coupling of the serotonin metabotropic receptors, 5‐HT1, and 5‐HT5 receptors generally couple to Gi proteins, which inhibit AC, 5‐HT2 receptors activate Gq‐proteins coupled to phospholipase C, and 5‐HT4, 5‐HT6, and 5‐HT7 receptors stimulate AC through activation of Gs (Barnes and Sharp, 1999; Raymond et al., 2001). The axon projections of serotonin‐containing neurons are diffuse and reach many different brain regions. Therefore, it is not surprising that serotonin affects AMPAR activity in many different cell types. The diversity of the cellular effects mediated by the various serotonin receptors also explains opposing effects on the activity of AMPAR. Glutamatergic synaptic transmission between primary afferent fibers and spinal dorsal horn neurons is subjected to modulation from supraspinal structures, including the rostroventral medulla (RVM) (Fields et al., 1991; Zhuo and Gebhart, 1992, 1997; Sandkuhler, 1996). Many nerve terminals projecting from this region toward the spinal cord release serotonin, and this neurotransmitter has a biphasic effect on synaptic transmission (Hori et al., 1996; Li and Zhuo, 1998; Li et al., 1999; Wang and Zhuo, 2002). In the spinal cord of young rats, low concentrations of serotonin (50mM), or stimulation with the 5‐HT2 agonist 2,5‐dimethoxy‐4‐iodoamphetamine (DOI), cause a long‐lasting synaptic enhancement, at least in part by activation of silent glutamatergic synapses. The recruitment of AMPAR to these ‘‘silent synapses’’ is thought to involve binding of the C‐terminal regions of GluR2/3 to postsynaptic PDZ proteins, such as GRIP/ABP, but not PICK1 (Li et al., 1999). These results support the idea that GluR2/3‐GRIP interaction may stabilize AMPAR in the postsynaptic plasma membrane. However, PKC activation is necessary and sufficient for synaptic potentiation induced by low concentrations of 5‐HT (Li et al., 1999), which contrasts with the role of the kinase in the internalization of AMPAR that is responsible for LTD in cerebellar Purkinje neurons (see > Section 3.1.2). In these cells, PKC‐mediated phosphorylation of GluR2 in Ser880 is thought to cause receptor internalization, due to a reduction in the interaction of GluR2 with GRIP. High concentrations of serotonin (50mM) decrease synaptic transmission in the spinal cord of young rats, and this effect is mimicked by the 5‐HT1A receptor agonist 8‐hydroxy‐2‐(di‐n‐propylamino)tetralin (8‐OH‐DPAT) (Hori et al., 1996; Li and Zhuo, 1998). In adult mice, serotonin (5–100mM) produces mainly inhibition of synaptic responses in dorsal horn, but coapplication of low doses of 5‐HT and forskolin, which activated AC, causes a long‐lasting enhancement of synaptic responses (Wang and Zhuo, 2002). Coapplication of 5‐HT and forskolin in ‘‘silent synapses’’ triggers the appearance of AMPA/kainate‐receptor mediated EPSPs, and this effect is independent of NMDA‐receptor activation (Wang and Zhuo, 2002). This indicates that there is a synergism between 5‐HT and Ca2þ‐dependent, calmodulin‐regulated AC (AC1, AC8) in the recruitment of functional AMPA responses in the adult mammalian spinal cord. Trigeminal motoneurons express functional NMDA and non‐NMDA‐receptors activated by glutamate released from mesencephalic nucleus of V afferents, and are involved in a variety of oral‐motor behaviors. Serotonin enhances AMPA‐ and NMDA‐mediated currents in guinea pig trigeminal motoneurons in brain stem slices, and this effect is mimicked by the 5‐HT2 receptor agonist DOI (Trueblood et al., 1996). In agreement with these findings, there is a dense serotonergic innervation of the trigeminal motor nucleus (Steinbusch, 1981) originating from the raphe nuclei (Fritschy et al., 1988), and trigeminal neurons are known to possess 5‐HT2 receptors (Kolta et al., 1993; Pompeiano et al., 1994). NMDA‐ and non‐NMDA‐receptors are involved in sensory transmission to the ventrobasal thalamus, a region that also receives projections from serotonergic brainstem neurons (Salt, 1987). Iontophoretic

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ejection of serotonin at low currents enhances neuronal activity induced by kainate, NMDA, and quisqualate in the rat ventrobasal thalamus in vivo (Eaton and Salt, 1989), but it remains to be determined which serotonin receptors are involved. The mechanism whereby serotonin 5‐HT1A receptors modulate AMPAR responses was extensively investigated in acutely dissociated prefrontal cortical (PFC) pyramidal neurons (Cai et al., 2002b). In these cells, resting CaMKII activity is thought to contribute to AMPAR activity by keeping the receptors at a highly phosphorylated state. Stimulation with serotonin, or with the 5‐HT1A receptor agonist 8‐OH‐DPAT, reduces the amplitude of AMPA‐mediated currents, due to the inhibition of PKA. The resulting decrease in the phosphorylation of PP‐1 regulatory peptides, and the concomitant activation of PP‐1, may act on AMPAR by reducing the autophosphorylation of CaMKII at Thr286, which is required for its Ca2þ‐ independent activity (Miller and Kennedy, 1986; Miller et al., 1988). The changes in the CaMKII phosphorylation of GluR1 subunits in PFC neurons, at Ser831, may underlie the effect of 5‐HT1A receptors on AMPA currents in these cells (Cai et al., 2002b). Accordingly, phosphorylation of GluR1 at Ser831 increases the apparent single‐channel conductance (Derkach et al., 1999). However, because phosphorylation of the same site drives synaptic incorporation of the receptors (Hayashi et al., 2000), it remains to be determined whether changes in AMPAR traffic account for the effect of 5‐HT1A receptors on AMPA‐mediated currents in PFC neurons. In the cerebellum, serotonin is released from fine beaded axons distributed in all layers of the cortical area and from large terminals typical of mossy fibers, which do not form classically defined synaptic contacts (Dieudonne, 2001). Iontophoretic application of serotonin in the adult rat in vivo antagonizes the increase in the firing frequency of cerebellar Purkinje cells induced by kainate or glutamate, and to a lower extent by AMPA (Netzeband et al., 1993). The effect of serotonin on glutamate‐induced neuronal excitability is mimicked by 8‐OH‐DPAT (Netzeband et al., 1993), an agonist of the 5‐HT1A receptors. However, it is surprising that the 5‐HT1A receptor mRNA is hardly detected in the molecular/Purkinje cell layer in adult rats (Miquel et al., 1994). The Purkinje cells receive excitatory inputs from granule cells and the inferior olivary CFs, and constitute the entire output of the cerebellar cortex (Dieudonne, 2001). Stimulation of slices from adult rat cerebellum with 8‐OH‐DPAT or with the 5‐HT2 receptor agonist DOI also inhibits AMPA‐evoked cGMP accumulation (Maura et al., 1995), but it remains to be determined whether this effect is due to inhibition of the activity of the glutamate receptors. In the Aplysia nervous system, 5‐HT released by interneurons facilitates synapses between sensory and motor neurons in the abdominal ganglion (Mackey et al., 1989). Facilitation of these sensorimotor synapses underlies in part the sensitization of the siphon‐withdrawal reflex (Glanzman et al., 1989). In isolated siphon motor neurons, the activation of G‐protein coupled 5‐HT receptors induces rapidly a long‐term facilitation of AMPA‐type responses, by a mechanism dependent on intracellular Ca2þ (Chitwood et al., 2001). This Ca2þ dependence suggests a role for phospholipase C‐coupled metabotropic receptors (Ap5‐ HTB2) that have been described in the Aplysia nervous system (Li et al., 1995). Although the mechanism underlying the 5‐HT‐induced facilitation of AMPA‐type responses was not yet elucidated, it is likely to involve an increase in the number of AMPAR in the motor neuron membrane (Chitwood et al., 2001). A long‐term effect (24h) of 5‐HT on the response mediated by AMPA‐type receptors was also observed in Aplysia motoneurons stimulated with excitatory amino acid agonists (Trudeau and Castellucci, 1995). This effect is dependent on new protein synthesis and, interestingly, is also inhibited by intracellular Ca2þ chelation with EGTA (Trudeau and Castellucci, 1995). Fluoxetine (Prozac), a widely prescribed medication for the treatment of depression, enhances serotonergic neurotransmission by inhibiting the 5‐HT reuptake. Acute administration of fluoxetine increases GluR1 phosphorylation at Ser831 and Ser845, in the frontal cortex, hippocampus, and striatum (Svenningsson et al., 2002). Chronic administration does not impair the ability of fluoxetine to induce the phosphorylation of GluR1 at Ser845, in a DARPP‐32‐dependent manner. In contrast, the effect of fluoxetine on GluR1 phosphorylation at Ser831 is lost in mice chronically exposed to the serotonin reuptake inhibitor (Svenningsson et al., 2002). In agreement with the role of DARPP‐32 in the regulation of GluR1 phosphorylation by fluoxetine, acute administration of the antidepressant increases phosphorylation of DARPP‐32 at the PKA site, Thr34, and at the casein kinase‐1 site, Ser137, and decreases phosphorylation at the cyclin‐dependent kinase 5 site, Thr75 (Svenningsson et al., 2002). These changes contribute to the

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inhibition of protein phosphatase‐1 by DARPP‐32, thereby affecting GluR1 phosphorylation, mainly at Ser845. The role of serotonin in the expression of AMPAR was examined in vivo, by injecting rats with the 5‐HT metabolite blocker para‐chorophenylalanine, for a week (Shutoh et al., 2000). Depletion of 5‐HT increased the number of binding sites for (S)‐[3H]a‐amino‐3‐hydroxy‐5‐methyl‐4‐isoxazole propionate ([3H]AMPA) in membranes isolated from the cerebral cortex, and, accordingly, Western blot analysis showed an increase in the immunoreactivity with antibodies against GluR2/3 and GluR2. A decrease in the GluR1 protein levels was observed under the same conditions (Shutoh et al., 2000), and the resulting increase in the relative abundance of GluR2 is likely to decrease Ca2þ permeability of AMPAR.

3.4

Adenosine Receptors

The neuromodulator adenosine acts through activation of four different metabotropic receptors, named A1, A2A, A2B, and A3 (Dunwiddie and Masino, 2001). The role of A2 adenosine receptors in the modulation of AMPAR was investigated in the CA1 region of rat transverse hippocampal slices. These receptors are coupled to the activation of AC and when stimulated with the agonist DPMA (N6‐[2‐(3,5‐dimethoxyphenyl)‐2‐(2‐methylphenyl)‐ethyl]adenosine) enhance synaptic transmission during low‐frequency test pulses (Kessey and Mogul, 1997). This effect was attributed to an increase in AMPAR‐mediated responses, but it may involve either an increase in the number of functional AMPAR at the synapse or a change in the phosphorylation state of the receptor (Kessey and Mogul, 1997). NMDA‐independent LTP is also induced in the CA1 region by the application of a tetanus during perfusion with DPMA and in the presence of a NMDA‐receptor antagonist. This potentiation is not observed after induction of LTP via multiple tetani, suggesting that the two forms of plasticity are convergent on a common mechanism (Kessey and Mogul, 1997).

3.5

Cholinergic Receptors

Acetylcholine acts through the activation of ionotropic (nicotinic) and metabotropic (muscarinic) receptors. Nicotinic receptors are ion‐permeable channels, whereas muscarinic receptors are coupled to G‐proteins and can be divided into two main categories: M1, M3, and M5 receptors, which stimulate preferentially the Gq family of G‐proteins, and the M2 and M4 receptors that preferentially couple to the Gi family of G‐proteins (Clementi et al., 2000; Nathanson, 2000). The muscarinic receptor agonist carbachol increases gradually the responses to iontophoretically applied AMPA in CA1 pyramidal cells (Auerbach and Segal, 1996). This slow response may be due to a translocation of receptors to the membrane, and contrasts with the immediate potentiation of the responses to NMDA, which may arise from the phosphorylation of plasma membrane associated receptors. Although the identity of the muscarinic receptors responsible for the potentiation of AMPA responses in the CA1 pyramidal cells was not yet investigated, they may belong to the M2 group, because these receptors also account for the carbachol‐evoked LTP in these hippocampal neurons (Auerbach and Segal, 1996). Chronic administration of nicotine, for 10 days, increased the NMDA‐ and AMPA‐evoked release of [3H] noradrenaline from isolated hippocampal nerve terminals, and NMDA‐evoked release of [3H]dopamine from striatal synaptosomes (Risso et al., 2004). However, it remains to be determined whether there is an upregulation in the amount of presynaptic NMDA‐receptors and AMPAR subunits under these conditions.

3.6

Adrenergic Receptors

Adrenergic receptors are sensitive to the catecholamines adrenaline (epinephrine) and noradrenaline (norepinephrine), which bind to the same receptors. This family of receptors is classified into three

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categories, a1, a2, and b, which are further subdivided into three classes (Barnes, 1995; Garcia‐Sainz et al., 1999). Activation of b1‐adrenergic receptors, coupled to the stimulation of AC, enable the induction of LTP during prolonged 5Hz synaptic stimulation in the hippocampal CA1 region (Thomas et al., 1996; Winder et al., 1999). A recent study using CA1 minislices showed that isoproterenol, an agonist of b‐adrenergic receptors, produces a long‐lasting increase in the phosphorylation of GluR1 at Ser845 (Vanhoose and Winder, 2003). As phosphorylation of this site is required for NMDA‐receptor dependent LTP (see earlier), this may constitute a common mechanism through which a heterosynaptic input regulates homosynaptic plasticity. Interestingly, although stimulation with NMDA also increases cAMP in CA1 minislices, to levels higher than those attained in the presence of isoproterenol, it decreases GluR1 phosphorylation on Ser845. These findings indicate that the cAMP/PKA signaling cascade activated by b1‐adrenergic receptors is preferentially located to induce the phosphorylation of GluR1 at Ser845 (Vanhoose and Winder, 2003). Phosphorylation of GluR1 on this site is required, in addition to CaMKII activation, for the recruitment of AMPAR to the synapse (Esteban et al., 2003), and this may contribute to the induction of LTP in the presence of b‐adrenergic receptor agonists.

3.7

Other GPCR

3.7.1 Opioid Receptors Three main opioid receptors were identified by molecular cloning techniques, and termed m‐, k‐, and d‐ opioid receptors. These receptors are all coupled to the activation of Gi proteins, but differ in the sensitivity to endogenous peptide ligands (Rogers and Peterson, 2003). Dynorphin, the endogenous ligand for the k‐receptors, decreases the amplitude and slows the kinetics of the AMPAR‐mediated currents in acutely isolated dorsal horn neurons from the Rexed’s laminae I–IV. This effect is mediated by Gi/Go proteins, sensitive to pertussis toxin and coupled to the inhibition of AC (Kolaj et al., 1995). In agreement with these findings, low concentrations of dynorphin decrease AMPAR‐ mediated primary afferent neurotransmission in the substantia gelatinosa of the young rat spinal cord. This effect is not observed in the presence of a PKA inhibitor, suggesting that it is also mediated by the inhibition of PKA (Randic et al., 1995). In addition to the effect of k‐receptors, m‐opioid receptors also depress AMPAR‐mediated currents in single spinal dorsal horn neurons, by a mechanism dependent on the activation of G‐proteins (Yoshimura and Jessell, 1990). The substantia gelatinosa (lamina 2) of the gray matter of the spinal cord dorsal horn is the preferential site of termination of primary afferent fibers sensitive to noxious stimuli (Kumazawa and Perl, 1978; Light and Perl, 1979; Sugiura et al., 1986, 1989; Yoshimura and Jessell, 1989, 1990).

3.7.2 Somatostatin Receptors Somatostatin exerts its physiological effects via a family of six GPCR (sst1, sst2A, sst2B, sst3, sst4, and sst5) (Csaba and Dournaud, 2001). Recent studies showed that activation of sst2 receptors, which interact with Gi and Go proteins (Csaba and Dournaud, 2001), decreases the AMPA component of glutamatergic synaptic responses in the hypothalamus, although they are without effect on pharmacologically isolated AMPA currents. The modulation of AMPA receptors by sst2 receptors depends on increases in the [Ca2þ]i, due to Ca2þ entry through NMDA‐receptors or calcium release from intracellular stores following activation of mGluRs, as shown in mediobasal hypothalamic cultures (Peineau et al., 2003). These results suggest that sst2 receptors may become phosphorylated/dephosphorylated following intracellular Ca2þ rise, and this may affect their ability to modulate the activity of AMPAR. Alternatively, sst2 receptors may act synergistically with the [Ca2þ]i rise in the modulation of the activity of a signaling molecule that changes the activity of AMPAR (Peineau et al., 2003).

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3.7.3 Vasopressin and Oxytocin Receptors Arginine‐vasopressin (AVP) modulates AMPA‐evoked currents in two populations of magnocellular neurons (MCNs) of the supraoptic nucleus (SON) (Hirasawa et al., 2003). These cells release AVP and oxytocin (OXT) within the SON from their dendrites and soma (Moos et al., 1984; Mason et al., 1986; Pow and Morris, 1989) to control their own electrical and secretory activity (Leng et al., 1999). In vasopressin neurons, AVP decreases AMPA‐induced currents through the activation of V1a receptors, whereas a facilitation of AMPA‐evoked currents by the agonist was observed in oxytocin neurons. The latter effects are mediated by oxytocin receptors (Hirasawa et al., 2003). V1a and oxytocin receptors are both coupled to the activation of phospholipase C via Gq family of G‐proteins (Michell et al., 1979; Lambert et al., 1994; Battle et al., 2000) and, therefore, differences in the signaling mechanisms are not expected to account for the differential effect on AMPA‐evoked currents in AVP and OXT neurons. These two neuronal populations possess functionally different non‐NMDA‐receptors, because of a differential combination of receptor subunits and/or their modulation (Stern et al., 1999). These differences may therefore explain the differential regulation of AMPA‐induced currents in the AVP and OXT neurons. The effect of AVP on non‐NMDA‐receptors indicates that when the peptide is released within the SON it favors an excitation of OXT neurons and inhibits AVP neurons. This indicates that the activity of AVP neurons would be slowed down following a high level of activity (Hirasawa et al., 2003).

3.7.4 Tachykinin Receptors The mammalian tachykinins include substance P, neurokinin A (NKA), and neurokinin B (NKB), which exert their biological effects by activation of three different receptors, named NK1, NK2, and NK3 (Severini et al., 2002; Pennefather et al., 2004). Substance P binds with high affinity to NK1, whereas NKA and NKB bind preferentially to NK2 and NK3, respectively. The tachykinin receptors belong to the GPCR family, and are coupled to the activation of phospholipase C (Khawaja and Rogers, 1996). Accordingly, in dorsal horn neurons, where the effect of tachykinins on AMPAR was investigated to some extent, substance P stimulates phosphoinositide turnover and mobilizes Ca2þ from internal stores (Mantyh et al., 1984; Womack et al., 1988). Pretreatment of dorsal horn neurons acutely isolated from the Rexed’s laminae I–III with substance P or NKA enhances the amplitude of the AMPA‐induced currents in a subpopulation of cells, and this effect is sensitive to [Ca2þ]i chelation with BAPTA (Rusin et al., 1992, 1993). In agreement with these findings GR 73632, a NK1‐ selective agonist, enhances the response of rat dorsal and ventral horn neurons to AMPA, as measured with extracellular electrodes implanted into the spinal cord (Cumberbatch et al., 1995). Furthermore, inhibition of NK1 receptors with selective antagonists reduces the responses to AMPA, suggesting that there is a tonic release of tachykinins that modulate AMPAR‐mediated glutamatergic neurotransmission in the spinal cord (Chizh et al., 1995). The NK3 receptor agonist senktide also enhances the response to AMPA in the spinal dorsal horn neurons (Cumberbatch et al., 1995). In the same study, the NK2 receptor agonist GR64349 was shown to decrease the response to AMPA and kainate (Cumberbatch et al., 1995), in contrast with the effect of NKA observed in a subpopulation of dorsal horn neurons acutely isolated from the Rexed’s laminae I–III (see earlier). However, the results showing an enhancement of the responses to kainate by the nonpeptide NK2 receptor antagonist GR159897 in the rat spinal cord (Chizh et al., 1995) suggest that these receptors increase the activity of non‐NMDA‐receptors. Because AMPAR are involved in mediating both nociceptive and non‐ nociceptive responses (Dougherty et al., 1992; Cumberbatch et al., 1994), alteration of their activity by tachykinins would be expected to affect spinal transmission in both cases.

3.7.5 Natriuretic Peptide Receptors A‐ and B‐type natriuretic peptide receptors, activated by atrial natriuretic peptide (ANP), brain natriuretic peptide (BNP), and C‐type natriuretic peptide (CNP), possess a guanylate cyclase domain that gives rise

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to cGMP when the receptors are activated by the ligand (Misono, 2002). The nucleus of the tractus solitarius (NTS) contains both ANP, derived from projections from the hypothalamus, and BNP, present in afferent projections to the NTS from the nodose ganglia (Standaert et al., 1986; Glaum and Miller, 1993). Exogenously applied BNP‐induced a long‐lasting potentiation of AMPA‐evoked currents in neurons of the dorsomedial subdivision of the NTS. Although the cell‐permeable analog of cGMP had a similar effect, the molecular mechanisms responsible for the potentiation of AMPA responses by BNP remain unknown (Glaum and Miller, 1993).

4

Regulation of AMPA Receptors by Receptor Tyrosine Kinases

4.1

Insulin and IGF Receptors

Insulin and its receptor are present in discrete regions throughout the brain, with the highest density found in the cerebral cortex and hippocampus (Hill et al., 1986; Kar et al., 1993; Wickelgren, 1998). Neurons synthesize and release insulin upon membrane depolarization (Clarke et al., 1986; Wozniak et al., 1993), and insulin receptors are highly expressed in neuronal cell bodies and synapses (Wozniak et al., 1993; Jonas et al., 1997; Abbott et al., 1999). These receptors have tyrosine kinase activity and activate several signaling pathways, including the phosphatidylinositol‐3‐kinase (PI3‐K) pathway, via the insulin receptor substrate proteins (IRS), and the mitogen‐activated protein kinase (MAPK) signaling pathway, through the adaptor proteins Shc and Gab‐1 (Saltiel and Pessin, 2002). Stimulation of cultured hippocampal neurons with insulin induces the internalization of GluR1‐ containing AMPAR (Beattie et al., 2000; Lin et al., 2000; Man et al., 2000; Zhou et al., 2001; Ahmadian et al., 2004), and studies using transfected HEK293 cells and hippocampal neurons showed that insulin stimulates AMPAR endocytosis via a mechanism requiring clathrin, dynamin and the last 15 amino acids in the C‐terminal region of the GluR2 subunit (Lin et al., 2000; Man et al., 2000; Ahmadian et al., 2004). Point mutation of the three tyrosine residues in this region eliminates insulin‐stimulated endocytosis of GluR2 (Ahmadian et al., 2004). The internalization of GluR1‐containing AMPAR in response to insulin follows a pattern distinct from that observed when internalization is induced by activating AMPA‐ or NMDA‐ receptors. Furthermore, there is a distinct dependence on extracellular Ca2þ and sensitivity to inhibitors of protein phosphatases (PP‐1 and protein phosphatase 2A, calcineurin and tyrosine phosphatase inhibitors) when internalization of AMPAR is induced by insulin or by activation of ionotropic glutamate receptors (Beattie et al., 2000; Lin et al., 2000). In addition, inhibition of tyrosine and serine/threonine kinases, with genistein and staurosporin, respectively, strongly inhibits the insulin‐induced internalization of AMPAR, yet does not block the AMPAR endocytosis caused by NMDA or AMPAR activation (Beattie et al., 2000). This indicates that the intracellular signaling mechanisms leading to AMPAR endocytosis in response to insulin are distinct from those triggered by NMDA and AMPA. Accordingly, GluR1 internalization elicited by insulin and AMPA treatments is additive in cultured hippocampal neurons (Beattie et al., 2000). The internalization of AMPAR in hippocampal neurons stimulated with insulin is associated with an increase in the rate of GluR1 insertion into the membrane, generating a new steady state in which internalization is in equilibrium with recycling to the surface (Lin et al., 2000; Passafaro et al., 2001). The exocytosis of GluR1‐induced by insulin is sensitive to the PI3‐K inhibitor wortmannin, but it remains to be determined how this signaling pathway activated by insulin receptors contributes to the translocation of the receptors (Hei, 1988; Passafaro et al., 2001; Claeys et al., 2002). As GluR1 may be delivered to nonsynaptic loci before lateral translocation into the synapse, this may lead to a reduction in the number of synaptic AMPAR. The effect of insulin on the internalization of AMPAR in hippocampal neurons is correlated with LTD‐ induced by insulin (insulin‐LTD) observed at the mossy fiber synapses and Shaffer collateral‐CA1 synapses, but not at associational commissural‐CA3 synapses (Huang et al., 2003; Ahmadian et al., 2004). Differences in the postsynaptic distribution of insulin receptors may account for the differential effect of insulin (Kar et al., 1993). Insulin‐LTD is expressed at the postsynaptic level, and involves activation of the PI3‐K pathway, in addition to a rise in the [Ca2þ]i, possibly due to Ca2þ entry through L‐type voltage‐gated

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channels. In agreement with the results reported in cultured neurons, a rapid clathrin‐mediated endocytosis of GluR1 was also observed in association with clathrin‐LTD formation in hippocampal slices (Huang et al., 2003). Phosphorylation of GluR2 on tyrosine at the C‐terminal region following activation of insulin receptors plays an important role in the internalization of the receptors in insulin‐LTD. In vitro and in vivo studies suggested that Src may be the tyrosine kinase responsible for phosphorylating GluR2 C‐terminal (Ahmadian et al., 2004). Insulin‐induced LTD and low‐frequency stimulation induced LTD are not mutually occlusive, suggesting that different mechanisms are involved. However, opposite results were reported for hippocampal CA1 neurons (Man et al., 2000). Interestingly, destruction of insulin receptors by intracerebroventricular injection of streptozotocin causes long‐term and progressive decreases in working and reference memory in the holeboard task and the passive avoidance paradigm, together with a permanent and ongoing cerebral energy deficit (Lannert and Hoyer, 1998). Insulin may also bind to the insulin‐like growth factor‐1 (IGF‐1) receptor, although the affinity of this receptor for IGF‐1 is about 100‐fold higher. Insulin and IGF‐1 receptors share a great level of homology, and seem to couple to the same signaling pathways (Kim and Accili, 2002). IGF‐1 depresses AMPA‐induced currents in cultured cerebellar Purkinje neurons, by a mechanism sensitive to inhibitors of clathrin‐ dependent endocytosis (Wang and Linden, 2000), resembling the mechanism for AMPAR internalization induced by insulin in the hippocampus (see earlier). However, IGF‐1 receptors do not contribute to LTD in cultures of Purkinje neurons, induced by stimulation with quisqualate/depolarization, or in granule cell‐ Purkinje neuron pairs, where presynaptic stimulation is paired with Purkinje neuron depolarization (Wang and Linden, 2000). The presence of IGF‐1 receptors in the cerebellum is well documented (Werther et al., 1990; Bondy et al., 1992; Garcia‐Segura et al., 1997; Sherrard et al., 1997). In addition to the short‐term effects on AMPA‐mediated synaptic transmission, insulin has a modulatory role in the functional maturation of the thalamocortical projection in culture (Plitzko et al., 2001). The proportion of functional glutamatergic synapses in thalamocortical cocultures increases during development in vitro, and this process is dramatically accelerated by chronic application of insulin (Plitzko et al., 2001). The effect of insulin is partially dependent on spontaneous activity, suggesting that insulin may facilitate the activity‐dependent conversion of silent synapses into functional ones. However, a general effect of insulin in promoting neuronal differentiation, including the increase in number and activity of synapses, may also account for its effects in differentiating neocortical neurons.

4.2

Neurotrophin Receptors

Neurotrophins are a family of secreted proteins that include nerve growth factor (NGF), brain‐derived neurotrophic factor (BDNF), neurotrophin‐3 (NT‐3), and neurotrophin 4/5 (NT‐4/5). The cellular effects of each neurotrophin are mediated through two different types of receptors, the Trk receptors and the p75 neurotrophin receptor. NGF binds preferentially to TrkA receptors, BDNF and NT‐4 to TrkB, and NT‐3 to TrkC receptors. Binding of neurotrophins to Trk receptors leads to receptor dimerization and autophosphorylation on intracellular tyrosine residues, and subsequent activation of intracellular signaling pathways, including the Ras/MAPK, phospholipase C‐g (PLC‐g), and PI3‐K/Akt (also called protein kinase B; Kaplan and Miller, 2000) pathways. Activation of these signaling pathways causes rapid‐ and long‐term alterations in the cell, the latter involving changes in transcriptional activity (Chao, 2003; Huang et al., 2003). The functional consequences of Trk receptor activation depend on the specific cell context and on the time course of neurotrophin availability. The p75 neurotrophin receptor binds all neurotrophins in addition to their precursors, called proneurotrophins. These receptors activate a set of signaling pathways distinct from those induced by Trk receptors, through adaptor proteins that bind to the cytoplasmic domain of p75. Activation of p75 receptors has been shown to mediate cell survival, cell death, and migration (Hempstead, 2002; Chao, 2003; Huang and Reichardt, 2003). Here, we will focus on the neurotrophin‐induced changes in the abundance of AMPAR, and on their activity and trafficking in and out of the synapse.

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4.2.1 Cultured Neocortical Neurons The studies concerning the effect of neurotrophins on AMPAR have been carried out using cultured neurons, mainly from the cerebral neocortex. Chronic exposure of developing neocortical neurons to BDNF increases GluR1 protein levels, particularly in the GABAergic population, expressing calbindin‐D and parvalbumin (Narisawa‐Saito et al., 1999a; Nagano et al., 2003). As BDNF does not affect GluR1 mRNA levels under the same conditions, the effect of the neurotrophin may be mediated by changes in the translational and/or posttranslational mechanisms (Narisawa‐Saito et al., 1999a). BDNF also upregulates GluR2/3 protein levels in developing neocortical neurons without a significant change in the mRNA for both receptor subunits. This contrasts with the reported induction of GluR2 promoter activity by BDNF and by glial‐derived neurotrophic factor (GDNF) in differentiated SH‐SY5Y cells, which appears to be mediated via a neuron‐restrictive silencer element (NRSE) present in the GluR2 promoter (Brene et al., 2000). Accordingly, in this cell line BDNF and GDNF increased GluR2 protein levels as determined by western blot (Brene et al., 2000). The upregulation in the total amount of GluR1 in BDNF‐stimulated cultured neocortical neurons is accompanied by an increase in the amount of the receptor subunit associated with the plasma membrane and by a marked increase in the inward membrane currents evoked by AMPA administration in the GABAergic neocortical neurons (Nagano et al., 2003). The increase in the AMPA currents is sensitive to K252a, a tyrosine kinase inhibitor, which also suppresses basal AMPAR activity, suggesting a role for the TrkB receptor in the effect of BDNF in cortical GABAergic neurons. Furthermore, chronic exposure of the cultures to BDNF increases the amplitude, but not the frequency, of spontaneous miniature excitatory postsynaptic currents (mEPSCs) in GABAergic neurons. Interestingly, the amplitude of mEPSC is significantly reduced in GABAergic neurons isolated from heterozygous BDNF‐knockout mice, suggesting that endogenous BDNF may contribute to the maintenance of AMPAR in these cells, even after development (Nagano et al., 2003). In contrast with the effects of BDNF, the neurotrophins NGF and NT‐3 do not affect AMPAR subunit protein levels in cultured neocortical neurons (Narisawa‐Saito et al., 1999a). To further understand the mechanism by which chronic exposure to BDNF increases GluR1 and GluR2/3 protein levels in cultured neocortical neurons, experiments were conducted using Fyn‐knockout mice (Narisawa‐Saito et al., 1999b). Fyn is a nonreceptor‐type PTK belonging to the Src‐family of PTKs, that is activated by TrkB receptors. These studies showed that activation of TrkB by BDNF enhances the protein levels of GluR1 and GluR2/3 via Fyn. However, other PTKs of the Src family may also contribute to the upregulation of AMPAR subunit protein levels by BDNF. Some of the molecular mechanisms involved in the translocation of AMPAR to the membrane following exposure to BDNF were investigated using cultured neocortical neurons as well as the heterologous system of human embryonic kidney (HEK) 293 cells carrying the TrkB receptor, and transiently transfected with GluR1 or GluR2 (Narisawa‐Saito et al., 2002). Stimulation of neocortical neurons with BDNF induces the incorporation of AMPAR into the membrane, by exocytosis, as detected by [3H]AMPA‐ binding studies using intact cells. The translocation of the receptors to the membrane occurs in a fairly rapid time scale of minutes to hours, and precedes the cellular increase in total AMPAR content. Although BDNF upregulates GluR1 and GluR2/3 protein levels associated with the membrane of cultured neurons, as determined by a biotinylation assay, studies with HEK‐TrkB cells expressing GluR1 or GluR2 showed an effect only for the latter subunit. The translocation of GluR2 to the membrane in BDNF‐stimulated cells involves the GluR2 NSF‐binding domain, but not the PDZ‐interaction site, in addition to Ca2þ influx. Interestingly, BDNF induces the translocation of GluR1 and GluR2 subunits to the membrane in HEK‐TrkB cells co‐transfected with both AMPAR subunits (Narisawa‐Saito et al., 2002). In addition to the effects on AMPAR protein levels, chronic stimulation with BDNF also enhances the abundance of the AMPAR interacting proteins SAP97, GRIP1, and PICK1 in neocortical neuronal cultures (Jourdi et al., 2003). Analysis of the mRNA levels under the same conditions showed an increase in gene expression for GRIP1 and PICK1, but not for SAP97. These observations suggest that BDNF may upregulate SAP97 protein levels by interfering directly with the synthesis of the protein. BDNF knockout

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mice also show reduced levels of SAP97, GRIP1, and PICK1 proteins, thus validating the results obtained in vitro studies (Jourdi et al., 2003). Immunoprecipitation studies using lysates from cultured neocortical neurons showed that chronic stimulation with BDNF increases the interaction between GluR1 and SAP97, as well as between GluR2 and GRIP1. In contrast, PICK1 association with GluR2/3 is not altered under the same conditions. The interaction with PDZ proteins plays an important role in the upregulation of AMPAR subunits by BDNF. Indeed, disruption of the interaction of GluR1 with SAP97, using Sindbis virus carrying the GluR1 C‐terminal decoy, counteracts the BDNF‐mediated increase in native GluR1, whereas the BDNF‐ dependent increase in GluR2/3 can still be detected. In addition, overexpression of the GluR2 C‐terminal decoy, which diminishes the interactions between GluR2/3 and GRIP1, decreases basal expression of GluR2/3 protein and blocks the BDNF‐dependent upregulation of GluR2/3. Interestingly, GluR2 C‐terminal decoy expression appears to counteract the BDNF‐triggered increase in native GluR1 as well (Jourdi et al., 2003). These findings support the hypothesis that the BDNF‐triggered increase in total AMPAR protein involves their molecular interaction via their carboxyl termini with PDZ proteins. Total lack of visual experience (dark rearing) is known to prolong the critical period and delay development of sensory functions in mammalian visual cortex (Timney et al., 1978; Fagiolini et al., 1994). Neurotrophins are known to modulate the development and synaptic efficacy of neurotransmitter systems that are affected by dark rearing, and the total lack of visual experience was shown to impair neurotrophin signaling (Viegi et al., 2002). Studies in dark‐reared rats showed that NGF is able to recover the induced downregulation on NR2A and PSD‐95 produced by total lack of visual experience, increasing the levels of both proteins. In contrast, exogenous administration of BDNF does not change the abundance of both proteins. This suggests that NGF is an elective trophic factor for the NMDA system in the visual cortex (Cotrufo et al., 2003). BDNF and NGF also produce an overshoot of GRIP protein levels, and NT4 also counteracts dark rearing‐induced downregulation of GRIP. This effect may arise from the activation of signaling pathways downstream to the TrkB receptor, shared by NT4 and BDNF (Chao, 2003; Huang et al., 2003). Both endogenous TrkB ligands could act on the same neurotransmitter system (i.e., AMPA) to modulate plastic modification of visual cortical circuitry during the critical period.

4.2.2 Cultured Hippocampal Neurons Long‐term treatment of hippocampal cultures with BDNF pontentiates both excitatory and inhibitory transmissions (McLean Bolton et al., 2000). Excitatory transmission is strengthened by BDNF, which increases the amplitude of miniature EPSCs in hippocampal cultures. This effect is due to an increase by BDNF of the quantal size of AMPAR‐mediated excitatory transmission, which does not require ongoing action potential activity. Other aspects of excitatory synaptic transmission in the same preparation are not substantially affected by the neurotrophin. However, BDNF also increases the frequency (but not the amplitude) of GABAA receptor‐mediated spontaneous quantal inhibitory transmission. These findings indicated that, although BDNF potentiates both excitatory and inhibitory synaptic transmission, in hippocampal cultures, it does so through distinct physiological mechanisms and, therefore, this neurotrophin can be seen as a mediator of activity‐dependent plasticity in vivo. The mechanisms underlying the effect of BDNF on excitatory neurotransmission in hippocampal cultures remain to be determined. The upregulation of synaptic activity by BDNF may be due to the insertion of AMPAR in the plasma membrane, thereby increasing the density of AMPAR at synapses (Carroll et al., 1999). Changes in the phosphorylation state of AMPA‐type glutamate receptors may also account for the effect of BDNF on excitatory synapses. Studies using autaptic cultures of hippocampal CA1 neurons also showed that chronic stimulation with BDNF induces long‐term enhancement of the strength of non‐NMDA‐receptor mediated synaptic currents, and this upregulation of mEPSC amplitude occurs via an increase in the size of unitary synaptic currents (Sherwood and Lo, 1999). Chronic BDNF treatment of the same preparation also decreases the degree of synaptic depression measured in response to paired stimuli (Sherwood and Lo, 1999). However, in contrast with long‐term treatment, brief BDNF applications affect neither the amplitude nor the extent of paired‐pulse depression of evoked EPSCs. Rather, BDNF acutely regulates the frequency of mEPSCs. Interestingly, very

Regulation of AMPA receptors by metabotropic receptors and receptor tyrosine kinases

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different effects of BDNF on quantal size at excitatory synapses were reported in mixed cultures of visual cortical neurons, where long‐term treatment with BDNF decrease, rather than increase, the amplitude of mEPSCs onto excitatory neurons (Rutherford et al., 1998). However, bipolar interneurons respond to BDNF similarly to hippocampal pyramidal neurons (Rutherford et al., 1998). Several experimental differences could account for this discrepancy in long‐term effects of BDNF, including brain region and cell‐type.

4.2.3 Nucleus Tractus Solitarius Neurons BDNF also markedly inhibits postsynaptic AMPAR‐mediated currents in a large subset of newborn nucleus tractus solitarius neurons (Balkowiec et al., 2000). This effect of BDNF is mimicked by NT4, but not by NGF, and blocked by the Trk tyrosine kinase inhibitor K252a, consistent with a requirement for TrkB receptor activation. Thus, there might be a close association between TrkB and AMPAR in these cells. One possible mechanism underlying acute inhibition of AMPA currents by BDNF is a rapid change in the number of available AMPAR, because BDNF signaling can influence the exocytosis of AMPAR (Narisawa‐ Saito et al., 2002). Alternatively, TrkB activation could lead to a change in AMPAR function without altering receptor availability.

4.3

Platelet‐Derived Growth Factor Receptors

Platelet derived growth factor (PDGF) interacts with target cells by binding with distinct specificities to two structurally related tyrosine kinase receptors, named a‐ and b‐receptors. As PDGF exists in solution as a dimer, it induces dimerization of the receptors followed by phosphorylation in trans between the two receptors in the complex. The tyrosine‐phosphorylated receptors interact with several different SH2‐ domain containing proteins, including effector enzymes (PLCg, PI3‐K, and members of the Src family) and adaptor proteins responsible for the activation of signaling cascades (Heldin and Westermark, 1999; Yu et al., 2003). Exposure of cultured neocortical neurons to PDGF for 5 days upregulates GluR1 and GluR2/3 protein levels via the Src‐family of PTKs (Narisawa‐Saito et al., 1999b). Accordingly, members of the Src family, including Fyn, interact and are activated by the PDGF receptor (Kypta et al., 1990; Hansen et al., 1997), and activation of Fyn mediates the effect of BDNF on the expression of AMPAR subunits (Narisawa‐Saito et al., 1999b, see earlier). However, the mechanisms underlying the effect of PDGF on the GluR1 and GluR2/3 protein levels, and the effect on AMPAR activity in neocortical neurons, remain to be determined. In contrast to the effect observed in cultured neurons, PDGF alone does not affect the expression of AMPAR subunits in cortical oligodendrocyte progenitor (O‐2A) cells (Chew et al., 1997).

4.4

Basic Fibroblast Growth Factor Receptors

Basic fibroblast growth factor (bFGF) binds to four plasma membrane associated tyrosine kinase receptors, with different affinities, but the mechanism of receptor activation is not yet fully elucidated. Autophosphorylation of the receptors upon ligand binding and dimerization allows the interaction and activation of PLCg, and binding to adaptor proteins that activate signaling cascades, including the Ras/MAPK pathway (Powers et al., 2000; Wang and McKeehan, 2003). Exposure of cultured hippocampal neurons to bFGF for 6–48h induces a time‐dependent and selective upregulation of GluR1 protein levels, but does not affect the abundance of GluR2/3 or GluR4 (Cheng et al., 1995). Under the same conditions, there is an increase in the [Ca2þ]i response to stimulation with AMPA in the cell bodies, suggesting that the newly synthesized receptors may be translocated to the membrane. bFGF treatment of cultured oligodendrocyte progenitor (O‐2A) cells for 3days also increases the steady‐state mRNA levels of GluR1 and GluR3 (Chew et al., 1997), but a 5‐day treatment is required to observe a

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significant induction of GluR4c mRNA (Gallo et al., 1994). Interestingly, although PDGF does not affect the expression of AMPAR subunits in O‐2A cells, it acts synergistically with bFGF to increase GluR1 mRNA levels as a result of increased transcriptional activity. In addition, bFGF increases GluR1 protein levels to some extent in O‐2A cells, and acts synergistically with PDGF in upregulating GluR1 protein levels and AMPAR activity (Chew et al., 1997).

5

Concluding Remarks

The molecular mechanisms underlying the modulation of AMPAR function by neurotransmitters, neuromodulators, and trophic factors will undoubtedly be further clarified in the next few years. These mechanisms contribute not only to transiently regulating AMPAR activity, which can be important in ensuring that at a certain time point a specific input has preferential influence on neuronal activity (e.g., the effects of dopamine in the NAc neurons), but also to adaptive brain function, as is exemplified by the contribution of mGluRs for LTD in cerebellum Purkinje cells, and the role of dopamine in controlling synaptic plasticity in the dorsal striatum and in the prefrontal cortex. Although most studies have so far focused on the regulation of AMPAR by a single type of metabotropic receptors or receptor tyrosine kinases, there may be also a cross‐talk between different neurotransmitter/neuromodulator systems in the regulation of AMPAR (e.g., effects of dopamine and mGluR in cultured retinal amacrine neurons). In those cases where such a mechanism exists, the regulation of AMPAR at a single moment will depend on the recent history of the synapse. Insights into the molecular mechanisms of regulation of AMPAR, together with more detailed knowledge about the neuronal circuits in which they operate, will provide the basis to further understand brain function and the molecular pathology of diseases of the glutamatergic synapse.

Acknowledgments The work in the authors’ laboratory was supported by FCT, FEDER, and Bissaya Barreto Foundation (Portugal).

References Abbott MA, Wells DG, Fallon JR. 1999. The insulin receptor tyrosine kinase substrate p58/53 and the insulin receptor are components of CNS synapses. J Neurosci 19: 7300-7308. Abraham WC, Bear MF. 1996. Metaplasticity: The plasticity of synaptic plasticity. Trends Neurosci 19: 126-130. Ahmadian G, Ju W, Liu L, Wyszynski M, Lee SH, et al. 2004. Tyrosine phosphorylation of GluR2 is required for insulin‐ stimulated AMPA receptor endocytosis and LTD. EMBO J 23: 1040-1050 [The authors show that insulin‐stimulated AMPAR endocytosis requires the phosphorylation of the GluR2 C‐terminal on tyrosine, possibly by Src. The effect of insulin on the phosphorylation of GluR2 appears to be necessary for insulin‐induced LTD at the Shaffer collateral CA1 synapses.]. Aiba A, Kano M, Chen C, Stanton ME, Fox GD, et al. 1994. Deficient cerebellar long‐term depression and impaired motor learning in mGluR1 mutant mice. Cell 79: 377-388. Alvarez FJ, Villalba RM, Carr PA, Grandes P, Somohano PM. 2000. Differential distribution of metabotropic glutamate

receptors 1a, 1b, and 5 in the rat spinal cord. J Comp Neurol 422: 464-487. Anneser JM, Borasio GD, Berthele A, Zieglgansberger W, Tolle TR. 1999. Differential expression of group I metabotropic glutamate receptors in rat spinal cord somatic and autonomic motoneurons: Possible implications for the pathogenesis of amyotrophic lateral sclerosis. Neurobiol Dis 6: 140-147. Anwyl R. 1999. Metabotropic glutamate receptors: Electrophysiological properties and role in plasticity. Brain Res Brain Res Rev 29: 83-120. Araki K, Meguro H, Kushiya E, Takayama C, Inoue Y, et al. 1993. Selective expression of the glutamate receptor channel delta 2 subunit in cerebellar Purkinje cells. Biochem Biophys Res Commun 197: 1267-1276. Ariano MA, Drago J, Sibley DR, Levine MS. 1998. Striatal excitatory amino acid receptor subunit expression in the D1A‐dopamine receptor‐deficient mouse. Dev Neurosci 20: 237-241.

Regulation of AMPA receptors by metabotropic receptors and receptor tyrosine kinases Ariano MA, Lewicki JA, Brandwein HJ, Murad F. 1982. Immunohistochemical localization of guanylate cyclase within neurons of rat brain. Proc Natl Acad Sci USA 79: 1316-1320. Ariano MA, Larson ER, Noblett KL, Sibley DR, Levine MS. 1997. Coexpression of striatal dopamine receptor subtypes and excitatory amino acid subunits. Synapse 26: 400-414. Armstrong N, Sun Y, Chen GQ, Gouaux E. 1998. Structure of a glutamate‐receptor ligand‐binding core in complex with kainate. Nature 395: 913-917. Auerbach JM, Segal M. 1996. Muscarinic receptors mediating depression and long‐term potentiation in rat hippocampus. J Physiol 492 (Pt 2): 479-493. Balkowiec A, Kunze DL, Katz DM. 2000. Brain‐derived neurotrophic factor acutely inhibits AMPA‐mediated currents in developing sensory relay neurons. J Neurosci 20: 1904-1911. Banke TG, Bowie D, Lee H, Huganir RL, Schousboe A, et al. 2000. Control of GluR1 AMPA receptor function by cAMP‐ dependent protein kinase. J Neurosci 20: 89-102. Bardo MT, Robinet PM, Mattingly BA, Margulies JE. 2001. Effect of 6‐hydroxydopamine or repeated amphetamine treatment on mesencephalic mRNA levels for AMPA glutamate receptor subunits in the rat. Neurosci Lett 302: 133-136. Barnes NM, Sharp T. 1999. A review of central 5‐HT receptors and their function. Neuropharmacology 38: 1083-1152. Barnes PJ. 1995. Beta‐adrenergic receptors and their regulation. Am J Respir Crit Care Med 152: 838-860. Barria A, Derkach V, Soderling T. 1997. Identification of the Ca2þ/calmodulin‐dependent protein kinase II regulatory phosphorylation site in the alpha‐amino‐3‐hydroxyl‐5‐ methyl‐4‐isoxazole‐propionate‐type glutamate receptor. J Biol Chem 272: 32727-32730. Battle T, Preisser L, Marteau V, Meduri G, Lambert M, et al. 2000. Vasopressin V1a receptor signaling in a rat choroid plexus cell line. Biochem Biophys Res Commun 275: 322-327. Baude A, Molnar E, Latawiec D, McIlhinney RA, Somogyi P. 1994. Synaptic and nonsynaptic localization of the GluR1 subunit of the AMPA‐type excitatory amino acid receptor in the rat cerebellum. J Neurosci 14: 2830-2843. Baude A, Nusser Z, Roberts JD, Mulvihill E, McIlhinney RA, et al. 1993. The metabotropic glutamate receptor (mGluR1 alpha) is concentrated at perisynaptic membrane of neuronal subpopulations as detected by immunogold reaction. Neuron 11: 771-787. Bear MF, Linden DJ. 2001. The mechanisms and meaning of long‐term synaptic depression in the mammalian brain. Synapses. Cowan WM, Sudhof TC, Stevens CF, editors. Baltimore, MD: The Johns Hopkins University Press; pp 455-517.

12

Beattie EC, Carroll RC, Yu X, Morishita W, Yasuda H, et al. 2000. Regulation of AMPA receptor endocytosis by a signaling mechanism shared with LTD. Nat Neurosci 3: 1291-1300. Benke TA, Luthi A, Isaac JT, Collingridge GL. 1998. Modulation of AMPA receptor unitary conductance by synaptic activity. Nature 393: 793-797. Bernard V, Somogyi P, Bolam JP. 1997. Cellular, subcellular, and subsynaptic distribution of AMPA‐type glutamate receptor subunits in the neostriatum of the rat. J Neurosci 17: 819-833. Betarbet R, Porter RH, Greenamyre JT. 2000. GluR1 glutamate receptor subunit is regulated differentially in the primate basal ganglia following nigrostriatal dopamine denervation. J Neurochem 74: 1166-1174. Bleakman D, Rusin KI, Chard PS, Glaum SR, Miller RJ. 1992. Metabotropic glutamate receptors potentiate ionotropic glutamate responses in the rat dorsal horn. Mol Pharmacol 42: 192-196. Bondy C, Werner H, Roberts CT Jr, Le Roith D. 1992. Cellular pattern of type‐I insulin‐like growth factor receptor gene expression during maturation of the rat brain: Comparison with insulin‐like growth factors I and II. Neuroscience 46: 909-923. Borgdorff AJ, Choquet D. 2002. Regulation of AMPA receptor lateral movements. Nature 417: 649-653. Bouyer JJ, Park DH, Joh TH, Pickel VM. 1984. Chemical and structural analysis of the relation between cortical inputs and tyrosine hydroxylase‐containing terminals in rat neostriatum. Brain Res 302: 267-275. Boxall AR, Garthwaite J. 1996. Long‐term depression in rat cerebellum requires both NO synthase and NO‐sensitive guanylyl cyclase. Eur J Neurosci 8: 2209-2212. Boyden ES, Katoh A, Raymond JL. 2004. Cerebellum‐dependent learning: The role of multiple plasticity mechanisms. Annu Rev Neurosci 27: 581-609. Brady AM, O’Donnell P. 2004. Dopaminergic modulation of prefrontal cortical input to nucleus accumbens neurons in vivo. J Neurosci 24: 1040-1049. Bredt DS, Nicoll RA. 2003. AMPA receptor trafficking at excitatory synapses. Neuron 40: 361-379. Brene S, Messer C, Nestler EJ. 1998. Expression of messenger RNAs encoding ionotropic glutamate receptors in rat brain: Regulation by haloperidol. Neuroscience 84: 813-823. Brene S, Messer C, Okado H, Hartley M, Heinemann SF, et al. 2000. Regulation of GluR2 promoter activity by neurotrophic factors via a neuron‐restrictive silencer element. Eur J Neurosci 12: 1525-1533. Brown JR, Arbuthnott GW. 1983. The electrophysiology of dopamine (D2) receptors: A study of the actions of dopamine on corticostriatal transmission. Neuroscience 10: 349-355.

309

310

12

Regulation of AMPA receptors by metabotropic receptors and receptor tyrosine kinases

Cai C, Coleman SK, Niemi K, Keinanen K. 2002a. Selective binding of synapse‐associated protein 97 to GluR‐A alpha‐ amino‐5‐hydroxy‐3‐methyl‐4‐isoxazole propionate receptor subunit is determined by a novel sequence motif. J Biol Chem 277: 31484-31490. Cai X, Gu Z, Zhong P, Ren Y, Yan Z. 2002b. Serotonin 5‐HT1A receptors regulate AMPA receptor channels through inhibiting Ca2þ/calmodulin‐dependent kinase II in prefrontal cortical pyramidal neurons. J Biol Chem 277: 36553-36562 [This paper shows that 5‐HT1A receptors decrease AMPA‐ mediated currents and the phosphorylation of GluR1 in prefrontal cortical neurons through inhibition of CaMKII, by inhibition of PKA and concomitant activation of protein phosphatase‐1.]. Calabresi P, Pisani AT, Mercuri NB, Bernardi G. 1992a. Long‐ term potentiation in the striatum is unmasked by removing the voltage‐dependent magnesium block of NMDA receptor channels. Eur J Neurosci 4: 929-935. Calabresi P, Maj R, Pisani A, Mercuri NB, Bernardi G. 1992b. Long‐term synaptic depression in the striatum: Physiological and pharmacological characterization. J Neurosci 12: 4224-4233. Calabresi P, Mercuri NB, Sancesario G, Bernardi G. 1993. Electrophysiology of dopamine‐denervated striatal neurons. Implications for Parkinson’s disease. Brain 116 (Pt 2): 433-452. Calabresi P, Gubellini P, Centonze D, Picconi B, Bernardi G, et al. 2000. Dopamine and cAMP‐regulated phosphoprotein 32 kDa controls both striatal long‐term depression and long‐term potentiation, opposing forms of synaptic plasticity. J Neurosci 20: 8443-8451. Calabresi P, Saiardi A, Pisani A, Baik JH, Centonze D, et al. 1997. Abnormal synaptic plasticity in the striatum of mice lacking dopamine D2 receptors. J Neurosci 17: 4536-4544. Campeau S, Davis M. 1995. Involvement of subcortical and cortical afferents to the lateral nucleus of the amygdala in fear conditioning measured with fear‐potentiated startle in rats trained concurrently with auditory and visual conditioned stimuli. J Neurosci 15: 2312-2327. Carlezon WA Jr, Nestler EJ. 2002. Elevated levels of GluR1 in the midbrain: A trigger for sensitization to drugs of abuse? Trends Neurosci 25: 610-615. Carroll RC, Lissin DV, von Zastrow M, Nicoll RA, Malenka RC. 1999. Rapid redistribution of glutamate receptors contributes to long‐term depression in hippocampal cultures. Nat Neurosci 2: 454-460. Carvalho AL, Duarte CB, Carvalho AP. 2000. Regulation of AMPA receptors by phosphorylation. Neurochem Res 25: 1245-1255. Carvalho AL, Kameyama K, Huganir RL. 1999. Characterization of phosphorylation sites on the glutamate receptor 4 subunit of the AMPA receptors. J Neurosci 19: 4748-4754.

Carvalho AL, Duarte CB, Faro CJ, Carvalho AP, Pires EV. 1998. Calcium influx through AMPA receptors and through calcium channels is regulated by protein kinase C in cultured retina amacrine‐like cells. J Neurochem 70: 2112-2119. Carvalho AL, Correia S, Faro CJ, Duarte CB, Carvalho AP, et al. 2002. Phosphorylation of GluR4 AMPA‐type glutamate receptor subunit by protein kinase C in cultured retina amacrine neurons. Eur J Neurosci 15: 465-474. Centonze D, Picconi B, Gubellini P, Bernardi G, Calabresi P. 2001. Dopaminergic control of synaptic plasticity in the dorsal striatum. Eur J Neurosci 13: 1071-1077. Centonze D, Gubellini P, Picconi B, Calabresi P, Giacomini P, et al. 1999. Unilateral dopamine denervation blocks corticostriatal LTP. J Neurophysiol 82: 3575-3579. Cepeda C, Buchwald NA, Levine MS. 1993. Neuromodulatory actions of dopamine in the neostriatum are dependent upon the excitatory amino acid receptor subtypes activated. Proc Natl Acad Sci USA 90: 9576-9580. Cerne R, Randic M. 1992. Modulation of AMPA and NMDA responses in rat spinal dorsal horn neurons by trans‐1‐ aminocyclopentane‐1,3‐dicarboxylic acid. Neurosci Lett 144: 180-184. Chao MV. 2003. Neurotrophins and their receptors: A convergence point for many signalling pathways. Nat Rev Neurosci 4: 299-309. Chao SZ, Ariano MA, Peterson DA, Wolf ME. 2002a. D1 dopamine receptor stimulation increases GluR1 surface expression in nucleus accumbens neurons. J Neurochem 83: 704-712. Chao SZ, Lu W, Lee HK, Huganir RL, Wolf ME. 2002b. D1 dopamine receptor stimulation increases GluR1 phosphorylation in postnatal nucleus accumbens cultures. J Neurochem 81: 984-992. Charpier S, Deniau JM. 1997. In vivo activity‐dependent plasticity at cortico‐striatal connections: Evidence for physiological long‐term potentiation. Proc Natl Acad Sci USA 94: 7036-7040. Chen L, Chetkovich DM, Petralia RS, Sweeney NT, Kawasaki Y, et al. 2000. Stargazin regulates synaptic targeting of AMPA receptors by two distinct mechanisms. Nature 408: 936-943. Cheng B, Furukawa K, O’Keefe JA, Goodman Y, Kihiko M, et al. 1995. Basic fibroblast growth factor selectively increases AMPA receptor subunit GluR1 protein level and differentially modulates Ca2þ responses to AMPA and NMDA in hippocampal neurons. J Neurochem 65: 2525-2536. Chew LJ, Fleck MW, Wright P, Scherer SE, Mayer ML, et al. 1997. Growth factor‐induced transcription of GluR1 increases functional AMPA receptor density in glial progenitor cells. J Neurosci 17: 227-240. Chitwood RA, Li Q, Glanzman DL. 2001. Serotonin facilitates AMPA‐type responses in isolated siphon motor neurons of

Regulation of AMPA receptors by metabotropic receptors and receptor tyrosine kinases Aplysia in culture. J Physiol 534: 501-510 [The authors show that 5‐HT increases the response of cultured Aplysia small siphon motor neurons to glutamate, the sensory neuron transmitter. It is suggested that the effect of 5‐HT is mediated by exocytotic insertion of glutamate receptors into the postsynaptic membrane during facilitation, because it was blocked by Botulinum toxin.]. Chizh BA, Cumberbatch MJ, Birch PJ, Headley PM. 1995. Endogenous modulation of excitatory amino acid responsiveness by tachykinin NK1 and NK2 receptors in the rat spinal cord. Br J Pharmacol 115: 1013-1019. Chung HJ, Steinberg JP, Huganir RL, Linden DJ. 2003. Requirement of AMPA receptor GluR2 phosphorylation for cerebellar long‐term depression. Science 300: 1751-1755. Chung HJ, Xia J, Scannevin RH, Zhang X, Huganir RL. 2000. Phosphorylation of the AMPA receptor subunit GluR2 differentially regulates its interaction with PDZ domain‐ containing proteins. J Neurosci 20: 7258-7267 [This paper shows that PKC phosphorylation of GluR2 on Ser880 plays an important role in the induction of cerebellar LTD.]. Churchill L, Swanson CJ, Urbina M, Kalivas PW. 1999. Repeated cocaine alters glutamate receptor subunit levels in the nucleus accumbens and ventral tegmental area of rats that develop behavioral sensitization. J Neurochem 72: 2397-2403. Claeys I, Simonet G, Poels J, Van Loy T, Vercammen L, et al. 2002. Insulin‐related peptides and their conserved signal transduction pathway. Peptides 23: 807-816. Clarke DW, Mudd L, Boyd FT Jr, Fields M, Raizada MK. 1986. Insulin is released from rat brain neuronal cells in culture. J Neurochem 47: 831-836. Clementi F, Fornasari D, Gotti C. 2000. Neuronal nicotinic receptors, important new players in brain function. Eur J Pharmacol 393: 3-10. Coleman SK, Cai C, Mottershead DG, Haapalahti JP, Keinanen K. 2003. Surface expression of GluR‐D AMPA receptor is dependent on an interaction between its C‐terminal domain and a 4.1 protein. J Neurosci 23: 798-806. Colledge M, Dean RA, Scott GK, Langeberg LK, Huganir RL, et al. 2000. Targeting of PKA to glutamate receptors through a MAGUK‐AKAP complex. Neuron 27: 107-119. Conn PJ, Pin JP. 1997. Pharmacology and functions of metabotropic glutamate receptors. Annu Rev Pharmacol Toxicol 37: 205-237. Conquet F, Bashir ZI, Davies CH, Daniel H, Ferraguti F, et al. 1994. Motor deficit and impairment of synaptic plasticity in mice lacking mGluR1. Nature 372: 237-243. Correia SS, Duarte CB, Faro CJ, Pires EV, Carvalho AL. 2003. Protein kinase C gamma associates directly with the GluR4 alpha‐amino‐3‐hydroxy‐5‐methyl‐4‐isoxazole propionate

12

receptor subunit. Effect on receptor phosphorylation. J Biol Chem 278: 6307-6313. Cotrufo T, Viegi A, Berardi N, Bozzi Y, Mascia L, et al. 2003. Effects of neurotrophins on synaptic protein expression in the visual cortex of dark‐reared rats. J Neurosci 23: 3566-3571. Crepel F, Audinat E, Daniel H, Hemart N, Jaillard D, et al. 1994. Cellular locus of the nitric oxide‐synthase involved in cerebellar long‐term depression induced by high external potassium concentration. Neuropharmacology 33: 13991405. Csaba Z, Dournaud P. 2001. Cellular biology of somatostatin receptors. Neuropeptides 35: 1-23. Cumberbatch MJ, Chizh BA, Headley PM. 1994. AMPA receptors have an equal role in spinal nociceptive and non‐nociceptive transmission. Neuroreport 5: 877-880. Cumberbatch MJ, Chizh BA, Headley PM. 1995. Modulation of excitatory amino acid responses by tachykinins and selective tachykinin receptor agonists in the rat spinal cord. Br J Pharmacol 115: 1005-1012. Dackis C, O’Brien C. 2003. Glutamatergic agents for cocaine dependence. Ann NY Acad Sci 1003: 328-345. David HN, Abraini JH. 2002. Group III metabotropic glutamate receptors and D1‐like and D2‐like dopamine receptors interact in the rat nucleus accumbens to influence locomotor activity. Eur J Neurosci 15: 869-875. Daw MI, Chittajallu R, Bortolotto ZA, Dev KK, Duprat F, et al. 2000. PDZ proteins interacting with C‐terminal GluR2/3 are involved in a PKC‐dependent regulation of AMPA receptors at hippocampal synapses. Neuron 28: 873-886. DeFrance JF, Sikes RW, Chronister RB. 1985. Dopamine action in the nucleus accumbens. J Neurophysiol 54: 1568-1577. Derkach V, Barria A, Soderling TR. 1999. Ca2þ/calmodulin‐ kinase II enhances channel conductance of alpha‐amino‐3‐ hydroxy‐5‐methyl‐4‐isoxazolepropionate type glutamate receptors. Proc Natl Acad Sci USA 96: 3269-3274. Descarries L, Watkins KC, Garcia S, Bosler O, Doucet G. 1996. Dual character, asynaptic and synaptic, of the dopamine innervation in adult rat neostriatum: A quantitative autoradiographic and immunocytochemical analysis. J Comp Neurol 375: 167-186. Detre JA, Nairn AC, Aswad DW, Greengard P. 1984. Localization in mammalian brain of G‐substrate, a specific substrate for guanosine 30 ,50 ‐cyclic monophosphate‐dependent protein kinase. J Neurosci 4: 2843-2849. Dieudonne S. 2001. Serotonergic neuromodulation in the cerebellar cortex: Cellular, synaptic, and molecular basis. Neuroscientist 7: 207-219. Dong H, O’Brien RJ, Fung ET, Lanahan AA, Worley PF, et al. 1997. GRIP: A synaptic PDZ domain‐containing protein that interacts with AMPA receptors. Nature 386: 279-284.

311

312

12

Regulation of AMPA receptors by metabotropic receptors and receptor tyrosine kinases

Dong Y, Saal D, Thomas M, Faust R, Bonci A, et al. 2004. Cocaine‐induced potentiation of synaptic strength in dopamine neurons: Behavioral correlates in GluRA(‐/‐) mice. Proc Natl Acad Sci USA 101: 14282-14287. Dougherty PM, Palecek J, Paleckova V, Sorkin LS, Willis WD. 1992. The role of NMDA and non‐NMDA excitatory amino acid receptors in the excitation of primate spinothalamic tract neurons by mechanical, chemical, thermal, and electrical stimuli. J Neurosci 12: 3025-3041. Dunwiddie TV, Masino SA. 2001. The role and regulation of adenosine in the central nervous system. Annu Rev Neurosci 24: 31-55. Eaton SA, Salt TE. 1989. Modulatory effects of serotonin on excitatory amino acid responses and sensory synaptic transmission in the ventrobasal thalamus. Neuroscience 33: 285-292. Ehlers MD. 2000. Reinsertion or degradation of AMPA receptors determined by activity‐dependent endocytic sorting. Neuron 28: 511-525. El‐Husseini AE, Schnell E, Chetkovich DM, Nicoll RA, Bredt DS. 2000. PSD‐95 involvement in maturation of excitatory synapses. Science 290: 1364-1368. El‐Husseini Ael D, Schnell E, Dakoji S, Sweeney N, Zhou Q, et al. 2002. Synaptic strength regulated by palmitate cycling on PSD‐95. Cell 108: 849-863. Endo S, Suzuki M, Sumi M, Nairn AC, Morita R, et al. 1999. Molecular identification of human G‐substrate, a possible downstream component of the cGMP‐dependent protein kinase cascade in cerebellar Purkinje cells. Proc Natl Acad Sci USA 96: 2467-2472. Esteban JA, Shi SH, Wilson C, Nuriya M, Huganir RL, et al. 2003. PKA phosphorylation of AMPA receptor subunits controls synaptic trafficking underlying plasticity. Nat Neurosci 6: 136-143. Eto M, Bock R, Brautigan DL, Linden DJ. 2002. Cerebellar long‐term synaptic depression requires PKC‐mediated activation of CPI‐17, a myosin/moesin phosphatase inhibitor. Neuron 36: 1145-1158. Fagiolini M, Pizzorusso T, Berardi N, Domenici L, Maffei L. 1994. Functional postnatal development of the rat primary visual cortex and the role of visual experience: Dark rearing and monocular deprivation. Vision Res 34: 709-720. Fan XD, Li XM, Juorio AV. 2000. Substantia nigra pars reticulata lesion induces preconvulsive behavior and changes in glutamate receptor gene expression in the rat brain. Brain Res 867: 40-51. Fan XD, Li XM, Ashe PC, Juorio AV. 1999. Lesion of the substantia nigra pars compacta downregulates striatal glutamate receptor subunit mRNA expression. Brain Res 850: 79-86. Feil R, Hartmann J, Luo C, Wolfsgruber W, Schilling K, et al. 2003. Impairment of LTD and cerebellar learning by

Purkinje cell‐specific ablation of cGMP‐dependent protein kinase I. J Cell Biol 163: 295-302 [The role of cGMP‐ dependent protein kinase type I in LTD and cerebellar learning was investigated in conditional knockout mice lacking the kinase selectively in Purkinje neurons. The results show that cGMP‐dependent protein kinase type I is required for cerebellar LTD and for adaptation of the vestibulo‐ocular reflex (VOR).]. Fields HL, Heinricher MM, Mason P. 1991. Neurotransmitters in nociceptive modulatory circuits. Annu Rev Neurosci 14: 219-245. Fitzgerald LW, Ortiz J, Hamedani AG, Nestler EJ. 1996. Drugs of abuse and stress increase the expression of GluR1 and NMDAR1 glutamate receptor subunits in the rat ventral tegmental area: Common adaptations among cross‐sensitizing agents. J Neurosci 16: 274-282 [The authors provide evidence that repeated cocaine or morphine treatment, as well as stress, elevates the levels of the GluR1 subunit of AMPAR in the ventral tegmental area (VTA) of the midbrain. This evidence, together with later studies, provides basis to the hypothesis that increased GluR1 expression in the VTA could be the trigger for sensitization to drugs of abuse (Carlezon and Nestler, 2002).]. Fitzgerald LW, Deutch AY, Gasic G, Heinemann SF, Nestler EJ. 1995. Regulation of cortical and subcortical glutamate receptor subunit expression by antipsychotic drugs. J Neurosci 15: 2453-2461. Floresco SB, Blaha CD, Yang CR, Phillips AG. 2001a. Dopamine D1 and NMDA receptors mediate potentiation of basolateral amygdala‐evoked firing of nucleus accumbens neurons. J Neurosci 21: 6370-6376. Floresco SB, Blaha CD, Yang CR, Phillips AG. 2001b. Modulation of hippocampal and amygdalar‐evoked activity of nucleus accumbens neurons by dopamine: Cellular mechanisms of input selection. J Neurosci 21: 2851-2860 [The authors assess the importance of dopamine in the integration of different limbic inputs to the nucleus accumbens. The study used extracellular single‐unit recordings of nucleus accumbens neurons in combination with sampling of dopamine efflux and the collected data suggest that increase in dopamine efflux by hippocampal afferents to the nucleus accumbens plays a critical role in the mechanism that ensures preferential reponse to the information conveyed from the hippocampus to the nucleus accumbens.]. Freund TF, Powell JF, Smith AD. 1984. Tyrosine hydroxylase‐ immunoreactive boutons in synaptic contact with identified striatonigral neurons, with particular reference to dendritic spines. Neuroscience 13: 1189-1215. Friedman E, Jin LQ, Cai GP, Hollon TR, Drago J, et al. 1997. D1‐like dopaminergic activation of phosphoinositide hydrolysis is independent of D1A dopamine

Regulation of AMPA receptors by metabotropic receptors and receptor tyrosine kinases receptors: Evidence from D1A knockout mice. Mol Pharmacol 51: 6-11. Fritschy JM, Lyons WE, Molliver ME, Grzanna R. 1988. Neurotoxic effects of p‐chloroamphetamine on the serotoninergic innervation of the trigeminal motor nucleus: A retrograde transport study. Brain Res 473: 261-270. Funabiki K, Mishina M, Hirano T. 1995. Retarded vestibular compensation in mutant mice deficient in delta 2 glutamate receptor subunit. Neuroreport 7: 189-192. Gallo V, Wright P, McKinnon RD. 1994. Expression and regulation of a glutamate receptor subunit by bFGF in oligodendrocyte progenitors. Glia 10: 149-153. Gallo V, Upson LM, Hayes WP, Vyklicky L Jr, Winters CA, et al. 1992. Molecular cloning and development analysis of a new glutamate receptor subunit isoform in cerebellum. J Neurosci 12: 1010-1023. Garcia‐Sainz JA, Vazquez‐Prado J, Villalobos‐Molina R. 1999. Alpha 1‐adrenoceptors: Subtypes, signaling, and roles in health and disease. Arch Med Res 30: 449-458. Garcia‐Segura LM, Rodriguez JR, Torres‐Aleman I. 1997. Localization of the insulin‐like growth factor I receptor in the cerebellum and hypothalamus of adult rats: An electron microscopic study. J Neurocytol 26: 479-490. Garris PA, Ciolkowski EL, Pastore P, Wightman RM. 1994. Efflux of dopamine from the synaptic cleft in the nucleus accumbens of the rat brain. J Neurosci 14: 6084-6093. Ghasemzadeh MB, Nelson LC, Lu XY, Kalivas PW. 1999. Neuroadaptations in ionotropic and metabotropic glutamate receptor mRNA produced by cocaine treatment. J Neurochem 72: 157-165. Glanzman DL, Mackey SL, Hawkins RD, Dyke AM, Lloyd PE, et al. 1989. Depletion of serotonin in the nervous system of Aplysia reduces the behavioral enhancement of gill withdrawal as well as the heterosynaptic facilitation produced by tail shock. J Neurosci 9: 4200-4213. Glaum SR, Miller RJ. 1993. Activation of metabotropic glutamate receptors produces reciprocal regulation of ionotropic glutamate and GABA responses in the nucleus of the tractus solitarius of the rat. J Neurosci 13: 1636-1641. Glaum SR, Slater NT, Rossi DJ, Miller RJ. 1992. Role of metabotropic glutamate (ACPD) receptors at the parallel fiber‐Purkinje cell synapse. J Neurophysiol 68: 1453-1462. Gomes AR, Correia SS, Carvalho AL, Duarte CB. 2003. Regulation of AMPA receptor activity, synaptic targeting and recycling: Role in synaptic plasticity. Neurochem Res 28: 1459-1473. Gomes AR, Cunha P, Nuriya M, Faro CJ, Huganir RL, et al. 2004. Metabotropic glutamate and dopamine receptors co‐regulate AMPA receptor activity through PKA in cultured chick retinal neurones: Effect on GluR4 phosphorylation and surface expression. J Neurochem 90: 673682 [In cultured retina amacrine cells, it is shown that

12

AMPAR are co‐regulated by metabotropic dopamine and glutamate receptors. The modulatory effects are due to GluR4 phosphorylation by PKA, which drives the surface expression of GluR4‐containing AMPAR.]. Gonon F. 1997. Prolonged and extrasynaptic excitatory action of dopamine mediated by D1 receptors in the rat striatum in vivo. J Neurosci 17: 5972-5978. Gonon F, Sundstrom L. 1996. Excitatory effects of dopamine released by impulse flow in the rat nucleus accumbens in vivo. Neuroscience 75: 13-18. Gonzalez‐Islas C, Hablitz JJ. 2003. Dopamine enhances EPSCs in layer II‐III pyramidal neurons in rat prefrontal cortex. J Neurosci 23: 867-875. Greengard P, Allen PB, Nairn AC. 1999. Beyond the dopamine receptor: The DARPP‐32/protein phosphatase‐1 cascade. Neuron 23: 435-447. Greengard P, Jen J, Nairn AC, Stevens CF. 1991. Enhancement of the glutamate response by cAMP‐dependent protein kinase in hippocampal neurons. Science 253: 1135-1138. Greger IH, Khatri L, Ziff EB. 2002. RNA editing at arg607 controls AMPA receptor exit from the endoplasmic reticulum. Neuron 34: 759-772. Groc L, Heine M, Cognet L, Brickley K, Stephenson FA, et al. 2004. Differential activity‐dependent regulation of the lateral mobilities of AMPA and NMDA receptors. Nat Neurosci 7: 695-696. Groenewegen HJ, Wright CI, Beijer AV, Voorn P. 1999. Convergence and segregation of ventral striatal inputs and outputs. Ann NY Acad Sci 877: 49-63. Gurden H, Takita M, Jay TM. 2000. Essential role of D1 but not D2 receptors in the NMDA receptor‐dependent long‐ term potentiation at hippocampal‐prefrontal cortex synapses in vivo. J Neurosci 20: RC106. Hall KU, Collins SP, Gamm DM, Massa E, DePaoli‐Roach AA, et al. 1999. Phosphorylation‐dependent inhibition of protein phosphatase‐1 by G‐substrate. A Purkinje cell substrate of the cyclic GMP‐dependent protein kinase. J Biol Chem 274: 3485-3495. Hansen K, Alonso G, Courtneidge SA, Ronnstrand L, Heldin CH. 1997. PDGF‐induced phosphorylation of Tyr28 in the N‐terminus of Fyn affects Fyn activation. Biochem Biophys Res Commun 241: 355-362. Hartell NA. 1994. Induction of cerebellar long‐term depression requires activation of glutamate metabotropic receptors. Neuroreport 5: 913-916. Harvey J, Lacey MG. 1996. Endogenous and exogenous dopamine depresses EPSCs in rat nucleus accumbens in vitro via D1 receptors activation. J Physiol 492 (Pt 1): 143-154. Harvey J, Lacey MG. 1997. A postsynaptic interaction between dopamine D1 and NMDA receptors promotes presynaptic inhibition in the rat nucleus accumbens via adenosine release. J Neurosci 17: 5271-5280.

313

314

12

Regulation of AMPA receptors by metabotropic receptors and receptor tyrosine kinases

Hayashi T, Huganir RL. 2004. Tyrosine phosphorylation and regulation of the AMPA receptor by SRC family tyrosine kinases. J Neurosci 24: 6152-6160. Hayashi Y, Shi SH, Esteban JA, Piccini A, Poncer JC, et al. 2000. Driving AMPA receptors into synapses by LTP and CaMKII: Requirement for GluR1 and PDZ domain interaction. Science 287: 2262-2267. Healy DJ, Meador‐Woodruff JH. 1997. Clozapine and haloperidol differentially affect AMPA and kainate receptor subunit mRNA levels in rat cortex and striatum. Brain Res Mol Brain Res 47: 331-338. Hei YJ. 1988. Recent progress in insulin signal transduction. J Pharmacol Toxicol Methods 40: 123-135. Heldin CH, Westermark B. 1999. Mechanism of action and in vivo role of platelet‐derived growth factor. Physiol Rev 79: 1283-1316. Hempstead BL. 2002. The many faces of p75NTR. Curr Opin Neurobiol 12: 260-267. Hidaka S, Totterdell S. 2001. Ultrastructural features of the nitric oxide synthase‐containing interneurons in the nucleus accumbens and their relationship with tyrosine hydroxylase‐containing terminals. J Comp Neurol 431: 139-154. Hill JM, Lesniak MA, Pert CB, Roth J. 1986. Autoradiographic localization of insulin receptors in rat brain: Prominence in olfactory and limbic areas. Neuroscience 17: 1127-1138. Hirasawa M, Mouginot D, Kozoriz MG, Kombian SB, Pittman QJ. 2003. Vasopressin differentially modulates non‐NMDA receptors in vasopressin and oxytocin neurons in the supraoptic nucleus. J Neurosci 23: 4270-4277. Hofmann F, Sold G. 1972. A protein kinase activity from rat cerebellum stimulated by guanosine‐30 :50 ‐monophosphate. Biochem Biophys Res Commun 49: 1100-1107. Hollmann M, Heinemann S. 1994. Cloned glutamate receptors. Annu Rev Neurosci 17: 31-108. Hori Y, Endo K, Takahashi T. 1996. Long‐lasting synaptic facilitation induced by serotonin in superficial dorsal horn neurones of the rat spinal cord. J Physiol 492 (Pt 3): 867-876. Hu G, Duffy P, Swanson C, Ghasemzadeh MB, Kalivas PW. 1999. The regulation of dopamine transmission by metabotropic glutamate receptors. J Pharmacol Exp Ther 289: 412-416. Huang EJ, Reichardt LF. 2003. Trk receptors: Roles in neuronal signal transduction. Annu Rev Biochem 72: 609-642. Huang CC, You JL, Lee CC, Hsu KS. 2003. Insulin induces a novel form of postsynaptic mossy fiber long‐term depression in the hippocampus. Mol Cell Neurosci 24: 831-841. Huang CC, You JL, Wu MY, Hsu KS. 2004. Rap1‐induced p38 mitogen‐activated protein kinase activation facilitates AMPA receptor trafficking via the GDI.Rab5 complex. Potential role in (S)‐3,5‐dihydroxyphenylglycene‐induced long term depression. J Biol Chem 279: 12286-12292 [The paper dissects some of the molecular events involved in the

endocytosis of AMPAR induced by group 1 metabotropic glutamate receptors at CA3‐CA1 synapses of the hippocampus. The loss of synaptic AMPAR is associated with a LTD of synaptic activity.]. Ito M. 1989. Long‐term depression. Annu Rev Neurosci 12: 85-102. Ito M. 2002. The molecular organization of cerebellar long‐ term depression. Nat Rev Neurosci 3: 896-902. Ito M, Karachot L. 1990. Messengers mediating long‐term desensitization in cerebellar Purkinje cells. Neuroreport 1: 129-132. Ito M, Karachot L. 1992. Protein kinases and phosphatase inhibitors mediating long‐term desensitization of glutamate receptors in cerebellar Purkinje cells. Neurosci Res 14: 27-38. Jiang ZG, North RA. 1991. Membrane properties and synaptic responses of rat striatal neurones in vitro. J Physiol 443: 533-553. Johnson LR, Aylward RL, Hussain Z, Totterdell S. 1994. Input from the amygdala to the rat nucleus accumbens: Its relationship with tyrosine hydroxylase immunoreactivity and identified neurons. Neuroscience 61: 851-865. Jonas EA, Knox RJ, Smith TC, Wayne NL, Connor JA, et al. 1997. Regulation by insulin of a unique neuronal Ca2þ pool and of neuropeptide secretion. Nature 385: 343-346. Jones KS, Van Dongen HM, Van Dongen AM. 2002. The NMDA receptor M3 segment is a conserved transduction element coupling ligand binding to channel opening. J Neurosci 22: 2044-2053. Jourdi H, Iwakura Y, Narisawa‐Saito M, Ibaraki K, Xiong H, et al. 2003. Brain‐derived neurotrophic factor signal enhances and maintains the expression of AMPA receptor‐associated PDZ proteins in developing cortical neurons. Dev Biol 263: 216-230. Kameyama K, Lee HK, Bear MF, Huganir RL. 1998. Involvement of a postsynaptic protein kinase A substrate in the expression of homosynaptic long‐term depression. Neuron 21: 1163-1175. Kano M, Kato M. 1987. Quisqualate receptors are specifically involved in cerebellar synaptic plasticity. Nature 325: 276-279. Kar S, Chabot JG, Quirion R. 1993. Quantitative autoradiographic localization of [125I]insulin‐like growth factor I, [125I]insulin‐like growth factor II, and [125I]insulin receptor binding sites in developing and adult rat brain. J Comp Neurol 333: 375-397. Kashiwabuchi N, Ikeda K, Araki K, Hirano T, Shibuki K, et al. 1995. Impairment of motor coordination, Purkinje cell synapse formation, and cerebellar long‐term depression in GluR delta 2 mutant mice. Cell 81: 245-252. Katoh A, Kitazawa H, Itohara S, Nagao S. 2000. Inhibition of nitric oxide synthesis and gene knockout of neuronal nitric

Regulation of AMPA receptors by metabotropic receptors and receptor tyrosine kinases oxide synthase impaired adaptation of mouse optokinetic response eye movements. Learn Mem 7: 220-226. Kerr JN, Wickens JR. 2001. Dopamine D‐1/D‐5 receptor activation is required for long‐term potentiation in the rat neostriatum in vitro. J Neurophysiol 85: 117-124. Kessey K, Mogul DJ. 1997. NMDA‐Independent LTP by adenosine A2 receptor‐mediated postsynaptic AMPA potentiation in hippocampus. J Neurophysiol 78: 19651972. Khawaja AM, Rogers DF. 1996. Tachykinins: Receptor to effector. Int J Biochem Cell Biol 28: 721-738. Kim JJ, Accili D. 2002. Signalling through IGF‐I and insulin receptors: Where is the specificity? Growth Horm IGF Res 12: 84-90. Kim JH, Vezina P. 1997. Activation of metabotropic glutamate receptors in the rat nucleus accumbens increases locomotor activity in a dopamine‐dependent manner. J Pharmacol Exp Ther 283: 962-968. Kim CH, Chung HJ, Lee HK, Huganir RL. 2001. Interaction of the AMPA receptor subunit GluR2/3 with PDZ domains regulates hippocampal long‐term depression. Proc Natl Acad Sci USA 98: 11725-11730. Kishimoto Y, Kawahara S, Suzuki M, Mori H, Mishina M, et al. 2001. Classical eyeblink conditioning in glutamate receptor subunit delta 2 mutant mice is impaired in the delay paradigm but not in the trace paradigm. Eur J Neurosci 13: 1249-1253. Knapp AG, Dowling JE. 1987. Dopamine enhances excitatory amino acid‐gated conductances in cultured retinal horizontal cells. Nature 325: 437-439 [This study provided the first evidence, which suggested that AMPA‐ and/or kainate‐ receptors could be regulated by phosphorylation. Using cultured horizontal retinal cells, the authors show that dopamine and cAMP enhance ionic conductances gated by kainate, and that the enhancing action of dopamine is specific to cone horizontal cells.]. Knapp AG, Schmidt KF, Dowling JE. 1990. Dopamine modulates the kinetics of ion channels gated by excitatory amino acids in retinal horizontal cells. Proc Natl Acad Sci USA 87: 767-771. Koenig JA. 2004. Signal reception: G protein‐coupled receptors. Molecular Biology of the Neuron. Davies RW, Morris BJ, editors. Oxford University Press; Oxford: pp. 215-248. Kohler M, Kornau HC, Seeburg PH. 1994. The organization of the gene for the functionally dominant alpha‐amino‐3‐ hydroxy‐5‐methylisoxazole‐4‐propionic acid receptor subunit GluR‐B. J Biol Chem 269: 17367-17370. Kolaj M, Cerne R, Randic M. 1995. The opioid peptide dynorphin modulates AMPA and kainate responses in acutely isolated neurons from the dorsal horn. Brain Res 671: 227-244.

12

Kolleker A, Zhu JJ, Schupp BJ, Qin Y, Mack V, et al. 2003. Glutamatergic plasticity by synaptic delivery of GluR‐ Blong‐containing AMPA receptors. Neuron 40: 1199-1212. Kolta A, Dubuc R, Lund JP. 1993. An immunocytochemical and autoradiographic investigation of the serotoninergic innervation of trigeminal mesencephalic and motor nuclei in the rabbit. Neuroscience 53: 1113-1126. Kombian SB, Malenka RC. 1994. Simultaneous LTP of non‐ NMDA‐ and LTD of NMDA‐receptor‐mediated responses in the nucleus accumbens. Nature 368: 242-246. Konnerth A, Dreessen J, Augustine GJ. 1992. Brief dendritic calcium signals initiate long‐lasting synaptic depression in cerebellar Purkinje cells. Proc Natl Acad Sci USA 89: 70517055. Kumazawa T, Perl ER. 1978. Excitation of marginal and substantia gelatinosa neurons in the primate spinal cord: Indications of their place in dorsal horn functional organization. J Comp Neurol 177: 417-434. Kypta RM, Goldberg Y, Ulug ET, Courtneidge SA. 1990. Association between the PDGF receptor and members of the src family of tyrosine kinases. Cell 62: 481-492. Lai SK, Tse YC, Yang MS, Wong CK, Chan YS, et al. 2003. Gene expression of glutamate receptors GluR1 and NR1 is differentially modulated in striatal neurons in rats after 6‐hydroxydopamine lesion. Neurochem Int 43: 639-653. Lambert RC, Dayanithi G, Moos FC, Richard P. 1994. A rise in the intracellular Ca2þ concentration of isolated rat supraoptic cells in response to oxytocin. J Physiol 478 (Pt 2): 275-287. Landsend AS, Amiry‐Moghaddam M, Matsubara A, Bergersen L, Usami S, et al. 1997. Differential localization of delta glutamate receptors in the rat cerebellum: Coexpression with AMPA receptors in parallel fiber‐spine synapses and absence from climbing fiber‐spine synapses. J Neurosci 17: 834-842. Lannert H, Hoyer S. 1998. Intracerebroventricular administration of streptozotocin causes long‐term diminutions in learning and memory abilities and in cerebral energy metabolism in adult rats. Behav Neurosci 112: 1199-1208. Laruelle M, Kegeles LS, Abi‐Dargham A. 2003. Glutamate, dopamine, and schizophrenia: From pathophysiology to treatment. Ann NY Acad Sci 1003: 138-158. Launey T, Endo S, Sakai R, Harano J, Ito M. 2004. Protein phosphatase 2A inhibition induces cerebellar long‐term depression and declustering of synaptic AMPA receptor. Proc Natl Acad Sci USA 101: 676-681. Le Doux JE, Cicchetti P, Xagoraris A, Romanski LM. 1990. The lateral amygdaloid nucleus: Sensory interface of the amygdala in fear conditioning. J Neurosci 10: 1062-1069. Lee HK, Kameyama K, Huganir RL, Bear MF. 1998. NMDA induces long‐term synaptic depression and dephosphorylation of the GluR1 subunit of AMPA receptors in hippocampus. Neuron 21: 1151-1162.

315

316

12

Regulation of AMPA receptors by metabotropic receptors and receptor tyrosine kinases

Lee HK, Barbarosie M, Kameyama K, Bear MF, Huganir RL. 2000. Regulation of distinct AMPA receptor phosphorylation sites during bidirectional synaptic plasticity. Nature 405: 955-959. Lee HK, Takamiya K, Han JS, Man H, Kim CH, et al. 2003. Phosphorylation of the AMPA receptor GluR1 subunit is required for synaptic plasticity and retention of spatial memory. Cell 112: 631-643. Lee SH, Simonetta A, Sheng M. 2004. Subunit rules governing the sorting of internalized AMPA receptors in hippocampal neurons. Neuron 43: 221-236. Lee SH, Liu L, Wang YT, Sheng M. 2002. Clathrin adaptor AP2 and NSF interact with overlapping sites of GluR2 and play distinct roles in AMPA receptor trafficking and hippocampal LTD. Neuron 36: 661-674. Leitges M, Kovac J, Plomann M, Linden DJ. 2004. A unique PDZ ligand in PKCalpha confers induction of cerebellar long‐term synaptic depression. Neuron 44: 585-594. Leng G, Brown CH, Russell JA. 1999. Physiological pathways regulating the activity of magnocellular neurosecretory cells. Prog Neurobiol 57: 625-655. Leonard AS, Davare MA, Horne MC, Garner CC, Hell JW. 1998. SAP97 is associated with the alpha‐amino‐3‐hydroxy‐ 5‐methylisoxazole‐4‐propionic acid receptor GluR1 subunit. J Biol Chem 273: 19518-19524. Letts VA, Felix R, Biddlecome GH, Arikkath J, Mahaffey CL, et al. 1998. The mouse stargazer gene encodes a neuronal Ca2þ‐channel gamma subunit. Nat Genet 19: 340-347. Levine MS, Li Z, Cepeda C, Cromwell HC, Altemus KL. 1996. Neuromodulatory actions of dopamine on synaptically‐ evoked neostriatal responses in slices. Synapse 24: 65-78. Lev‐Ram V, Nebyelul Z, Ellisman MH, Huang PL, Tsien RY. 1997. Absence of cerebellar long‐term depression in mice lacking neuronal nitric oxide synthase. Learn Mem 4: 169-177. Li P, Zhuo M. 1998. Silent glutamatergic synapses and nociception in mammalian spinal cord. Nature 393: 695-698. Li P, Kerchner GA, Sala C, Wei F, Huettner JE, et al. 1999. AMPA receptor‐PDZ interactions in facilitation of spinal sensory synapses. Nat Neurosci 2: 972-977 [The role of GluR2/3‐PDZ interactions in 5‐HT‐induced activation of silent synapses in the spinal cord was studied using synthetic peptides corresponding to the C‐terminal region of the two AMPAR subunits, which differentially affect the interaction with intracellular proteins. The results implicate AMPAR–PDZ interactions in sensory synaptic potentiation in young rats, and show that PKC is necessary and sufficient to activate silent synapses in the spinal cord.]. Li XC, Giot JF, Kuhl D, Hen R, Kandel ER. 1995. Cloning and characterization of two related serotonergic receptors from

the brain and the reproductive system of Aplysia that activate phospholipase C. J Neurosci 15: 7585-7591. Light AR, Perl ER. 1979. Spinal termination of functionally identified primary afferent neurons with slowly conducting myelinated fibers. J Comp Neurol 186: 133-150. Lin JW, Ju W, Foster K, Lee SH, Ahmadian G, et al. 2000. Distinct molecular mechanisms and divergent endocytotic pathways of AMPA receptor internalization. Nat Neurosci 3: 1282-1290. Lin JY, Dubey R, Funk GD, Lipski J. 2003. Receptor subtype‐ specific modulation by dopamine of glutamatergic responses in striatal medium spiny neurons. Brain Res 959: 251-262. Linden DJ, Connor JA. 1991. Participation of postsynaptic PKC in cerebellar long‐term depression in culture. Science 254: 1656-1659. Linden DJ, Smeyne M, Connor JA. 1993. Induction of cerebellar long‐term depression in culture requires postsynaptic action of sodium ions. Neuron 11: 1093-1100. Linden DJ, Dickinson MH, Smeyne M, Connor JA. 1991. A long‐term depression of AMPA currents in cultured cerebellar Purkinje neurons. Neuron 7: 81-89. Lohmann SM, Walter U, Miller PE, Greengard P, De Camilli P. 1981. Immunohistochemical localization of cyclic GMP‐ dependent protein kinase in mammalian brain. Proc Natl Acad Sci USA 78: 653-657. Lomeli H, Sprengel R, Laurie DJ, Kohr G, Herb A, et al. 1993. The rat delta‐1 and delta‐2 subunits extend the excitatory amino acid receptor family. FEBS Lett 315: 318-322. Lomeli H, Mosbacher J, Melcher T, Hoger T, Geiger JR, et al. 1994. Control of kinetic properties of AMPA receptor channels by nuclear RNA editing. Science 266: 1709-1713. Lu W, Monteggia LM, Wolf ME. 2002. Repeated administration of amphetamine or cocaine does not alter AMPA receptor subunit expression in the rat midbrain. Neuropsychopharmacology 26: 1-13. Lu W, Chen H, Xue CJ, Wolf ME. 1997. Repeated amphetamine administration alters the expression of mRNA for AMPA receptor subunits in rat nucleus accumbens and prefrontal cortex. Synapse 26: 269-280. Mackey SL, Kandel ER, Hawkins RD. 1989. Identified serotonergic neurons LCB1 and RCB1 in the cerebral ganglia of Aplysia produce presynaptic facilitation of siphon sensory neurons. J Neurosci 9: 4227-4235. Maguire G, Werblin F. 1994. Dopamine enhances a glutamate‐ gated ionic current in OFF bipolar cells of the tiger salamander retina. J Neurosci 14: 6094-6101. Mahan LC, Burch RM, Monsma FJ Jr, Sibley DR. 1990. Expression of striatal D1 dopamine receptors coupled to inositol phosphate production and Ca2þ mobilization in Xenopus oocytes. Proc Natl Acad Sci USA 87: 2196-2200.

Regulation of AMPA receptors by metabotropic receptors and receptor tyrosine kinases Malenka RC, Bear MF. 2004. LTP and LTD: An embarrassment of riches. Neuron 44: 5-21 [This review covers the currently established cellular and molecular mechanisms underlying LTP and LTD, and discusses examples in which attempts have been made to address the functional roles of LTP and LTD, by examining whether synaptic modifications analogous to LTP or LTD actually occur in vivo in response to experience.]. Malenka RC, Kocsis JD. 1988. Presynaptic actions of carbachol and adenosine on corticostriatal synaptic transmission studied in vitro. J Neurosci 8: 3750-3756. Malinow R, Malenka RC. 2002. AMPA receptor trafficking and synaptic plasticity. Annu Rev Neurosci 25: 103-126 [The authors review the growing literature that supports a critical role for AMPA receptor trafficking in LTP and LTD, and focus on the different roles proposed for specific AMPA receptor subunits and their interaction partners.]. Mammen AL, Kameyama K, Roche KW, Huganir RL. 1997. Phosphorylation of the alpha‐amino‐3‐hydroxy‐5‐methylisoxazole4‐propionic acid receptor GluR1 subunit by calcium/calmodulin‐dependent kinase II. J Biol Chem 272: 32528-32533. Man HY, Lin JW, Ju WH, Ahmadian G, Liu L, et al. 2000. Regulation of AMPA receptor‐mediated synaptic transmission by clathrin‐dependent receptor internalization. Neuron 25: 649-662 [The authors show that the activation of postsynaptic insulin receptors cause a LTD of synaptic transmission in the CA1 region of the hippocampus, by a mechanism overlapping with CA1 LTD induced by low‐ frequency stimulation. Studies using HEK293 cells and neurons indicate that insulin stimulates AMPAR endocytosis, by a mechanism dependent on clathrin, dynamin, and the C‐terminal region of GluR2.]. Mangiavacchi S, Wolf ME. 2004. D1 dopamine receptor stimulation increases the rate of AMPA receptor insertion onto the surface of cultured nucleus accumbens neurons through a pathway dependent on protein kinase A. J Neurochem 88: 1261-1271. Mantyh PW, Pinnock RD, Downes CP, Goedert M, Hunt SP. 1984. Correlation between inositol phospholipid hydrolysis and substance P receptors in rat CNS. Nature 309: 795-797. Manzoni O, Michel JM, Bockaert J. 1997. Metabotropic glutamate receptors in the rat nucleus accumbens. Eur J Neurosci 9: 1514-1523. Masland RH. 2001. The fundamental plan of the retina. Nat Neurosci 4: 877-886. Mason WT, Hatton GI, Ho YW, Chapman C, Robinson IC. 1986. Central release of oxytocin, vasopressin and neurophysin by magnocellular neurone depolarization: Evidence in slices of guinea pig and rat hypothalamus. Neuroendocrinology 42: 311-322.

12

Matsuda S, Mikawa S, Hirai H. 1999. Phosphorylation of serine‐880 in GluR2 by protein kinase C prevents its C terminus from binding with glutamate receptor‐interacting protein. J Neurochem 73: 1765-1768. Matsuda S, Launey T, Mikawa S, Hirai H. 2000. Disruption of AMPA receptor GluR2 clusters following long‐term depression induction in cerebellar Purkinje neurons. EMBO J 19: 2765-2774 [Stimulations inducing LTD in cultured cerebellar Purkinje cells are shown to cause phosphorylation of GluR2 on Ser880. This phosphorylation was accompanied by a reduction in the interaction with GRIP, a disruption of postsynaptic GluR2 clusters and internalization of the receptors.]. Maura G, Guadagnin A, Raiteri M. 1995. Low nanomolar serotonin inhibits the glutamate receptor/nitric oxide/ cyclic GMP pathway in slices from adult rat cerebellum. Neuroscience 68: 455-463. McDonald BJ, Chung HJ, Huganir RL. 2001. Identification of protein kinase C phosphorylation sites within the AMPA receptor GluR2 subunit. Neuropharmacology 41: 672-679. McGlade‐McCulloh E, Yamamoto H, Tan SE, Brickey DA, Soderling TR. 1993. Phosphorylation and regulation of glutamate receptors by calcium/calmodulin‐dependent protein kinase II. Nature 362: 640-642. McLean Bolton M, Pittman AJ, Lo DC. 2000. Brain‐derived neurotrophic factor differentially regulates excitatory and inhibitory synaptic transmission in hippocampal cultures. J Neurosci 20: 3221-3232. Meador‐Woodruff JH, Healy DJ. 2000. Glutamate receptor expression in schizophrenic brain. Brain Res Brain Res Rev 31: 288-294. Mi R, Sia GM, Rosen K, Tang X, Moghekar A, et al. 2004. AMPA receptor‐dependent clustering of synaptic NMDA receptors is mediated by stargazin and NR2A/B in spinal neurons and hippocampal interneurons. Neuron 44: 335-349. Mi R, Tang X, Sutter R, Xu D, Worley P, et al. 2002. Differing mechanisms for glutamate receptor aggregation on dendritic spines and shafts in cultured hippocampal neurons. J Neurosci 22: 7606-7616. Michell RH, Kirk CJ, Billah MM. 1979. Hormonal stimulation of phosphatidylinositol breakdown with particular reference to the hepatic effects of vasopressin. Biochem Soc Trans 7: 861-865. Miller SG, Kennedy MB. 1986. Regulation of brain type II Ca2þ/calmodulin‐dependent protein kinase by autophosphorylation: A Ca2þ‐triggered molecular switch. Cell 44: 861-870. Miller SG, Patton BL, Kennedy MB. 1988. Sequences of autophosphorylation sites in neuronal type II CaM kinase that control Ca2þ‐independent activity. Neuron 1: 593-604.

317

318

12

Regulation of AMPA receptors by metabotropic receptors and receptor tyrosine kinases

Miquel MC, Kia HK, Boni C, Doucet E, Daval G, et al. 1994. Postnatal development and localization of 5‐HT1A receptor mRNA in rat forebrain and cerebellum. Brain Res Dev Brain Res 80: 149-157. Misono KS. 2002. Natriuretic peptide receptor: Structure and signaling. Mol Cell Biochem 230: 49-60. Missale C, Nash SR, Robinson SW, Jaber M, Caron MG. 1998. Dopamine receptors: From structure to function. Physiol Rev 78: 189-225. Moghaddam B, Gruen RJ, Roth RH, Bunney BS, Adams RN. 1990. Effect of L‐glutamate on the release of striatal dopamine: In vivo dialysis and electrochemical studies. Brain Res 518: 55-60. Moos F, Freund‐Mercier MJ, Guerne Y, Guerne JM, Stoeckel ME, et al. 1984. Release of oxytocin and vasopressin by magnocellular nuclei in vitro: Specific facilitatory effect of oxytocin on its own release. J Endocrinol 102: 63-72. Morari M, Marti M, Sbrenna S, Fuxe K, Bianchi C, et al. 1998. Reciprocal dopamine‐glutamate modulation of release in the basal ganglia. Neurochem Int 33: 383-397. Mori‐Okamoto J, Okamoto K, Tatsuno J. 1993. Intracellular mechanisms underlying the suppression of AMPA responses by trans‐ACPD in cultured chick Purkinje neurons. Mol Cell Neurosci 4: 375-386. Mosbacher J, Schoepfer R, Monyer H, Burnashev N, Seeburg PH, et al. 1994. A molecular determinant for submillisecond desensitization in glutamate receptors. Science 266: 1059-1062. Muresan Z, Besharse JC. 1993. D2‐like dopamine receptors in amphibian retina: Localization with fluorescent ligands. J Comp Neurol 331: 149-160. Nagano T, Yanagawa Y, Obata K, Narisawa‐Saito M, Namba H, et al. 2003. Brain‐derived neurotrophic factor upregulates and maintains AMPA receptor currents in neocortical GABAergic neurons. Mol Cell Neurosci 24: 340-356. Nakagawa T, Futai K, Lashuel HA, Lo I, Okamoto K, et al. 2004. Quaternary structure, protein dynamics, and synaptic function of SAP97 controlled by L27 domain interactions. Neuron 44: 453-467. Nakanishi S, Maeda N, Mikoshiba K. 1991. Immunohistochemical localization of an inositol 1,4,5‐trisphosphate receptor, P400, in neural tissue: Studies in developing and adult mouse brain. J Neurosci 11: 2075-2086. Narisawa‐Saito M, Carnahan J, Araki K, Yamaguchi T, Nawa H. 1999a. Brain‐derived neurotrophic factor regulates the expression of AMPA receptor proteins in neocortical neurons. Neuroscience 88: 1009-1014. Narisawa‐Saito M, Iwakura Y, Kawamura M, Araki K, Kozaki S, et al. 2002. Brain‐derived neurotrophic factor regulates surface expression of alpha‐amino‐3‐hydroxy‐5‐methyl‐4‐ isoxazoleproprionic acid receptors by enhancing the N‐ethylmaleimide‐sensitive factor/GluR2 interaction in

developing neocortical neurons. J Biol Chem 277: 4090140910 [The effect of the neurotrophin BDNF on the translocation of AMPAR to the membrane is described in cultured neocortical neurons. This process requires intracellular Ca2þ and is sensitive to chemical and toxin inhibitors of exocytosis. Studies in HEK293 cells carrying the BDNF receptor TrkB and expressing GluR2 showed that the translocation of this receptor subunit to the membrane requires its NSF‐binding domain.]. Narisawa‐Saito M, Silva AJ, Yamaguchi T, Hayashi T, Yamamoto T, et al. 1999b. Growth factor‐mediated Fyn signaling regulates alpha‐amino‐3‐hydroxy‐5‐methyl‐4‐isoxazolepropionic acid (AMPA) receptor expression in rodent neocortical neurons. Proc Natl Acad Sci USA 96: 2461-2466 [The role of Src‐family protein tyrosine kinases in the upregulation of GluR1 and GluR2/3 protein levels by BDNF was shown in cultured neocortical neurons, using chemical inhibitors. The use of Fyn‐knockout mice showed a role for this kinase in the effect of BDNF on AMPAR protein levels.]. Nathanson NM. 2000. A multiplicity of muscarinic mechanisms: Enough signaling pathways to take your breath away. Proc Natl Acad Sci USA 97: 6245-6247. Nestler EJ. 2004. Historical review: Molecular and cellular mechanisms of opiate and cocaine addiction. Trends Pharmacol Sci 25: 210-218. Netzeband JG, Weathers LB, Strahlendorf HK, Strahlendorf JC. 1993. Serotonin depresses excitatory amino acid‐ induced excitation of cerebellar Purkinje cells in the adult rat in vivo. Brain Res 608: 145-149. Nguyen‐Legros J, Simon A, Caille I, Bloch B. 1997. Immunocytochemical localization of dopamine D1 receptors in the retina of mammals. Vis Neurosci 14: 545-551. Nicola SM, Malenka RC. 1997. Dopamine depresses excitatory and inhibitory synaptic transmission by distinct mechanisms in the nucleus accumbens. J Neurosci 17: 5697-5710. Nicola SM, Malenka RC. 1998. Modulation of synaptic transmission by dopamine and norepinephrine in ventral but not dorsal striatum. J Neurophysiol 79: 1768-1776. Nicola SM, Kombian SB, Malenka RC. 1996. Psychostimulants depress excitatory synaptic transmission in the nucleus accumbens via presynaptic D1‐like dopamine receptors. J Neurosci 16: 1591-1604. Nicola SM, Surmeier J, Malenka RC. 2000. Dopaminergic modulation of neuronal excitability in the striatum and nucleus accumbens. Annu Rev Neurosci 23: 185-215. Nishimune A, Isaac JT, Molnar E, Noel J, Nash SR, et al. 1998. NSF binding to GluR2 regulates synaptic transmission. Neuron 21: 87-97. O’Brien R, Xu D, Mi R, Tang X, Hopf C, et al. 2002. Synaptically targeted narp plays an essential role in the aggregation of AMPA receptors at excitatory synapses in cultured spinal neurons. J Neurosci 22: 4487-4498.

Regulation of AMPA receptors by metabotropic receptors and receptor tyrosine kinases O’Brien RJ, Xu D, Petralia RS, Steward O, Huganir RL, et al. 1999. Synaptic clustering of AMPA receptors by the extracellular immediate‐early gene product Narp. Neuron 23: 309-323. Ohishi H, Neki A, Mizuno N. 1998. Distribution of a metabotropic glutamate receptor, mGluR2, in the central nervous system of the rat and mouse: An immunohistochemical study with a monoclonal antibody. Neurosci Res 30: 65-82. Ohishi H, Akazawa C, Shigemoto R, Nakanishi S, Mizuno N. 1995. Distributions of the mRNAs for L‐2‐amino‐4‐phosphonobutyrate‐sensitive metabotropic glutamate receptors, mGluR4 and mGluR7, in the rat brain. J Comp Neurol 360: 555-570. Ohno M, Watanabe S. 1995. Persistent increase in dopamine release following activation of metabotropic glutamate receptors in the rat nucleus accumbens. Neurosci Lett 200: 113-116. Olveczky BP, Baccus SA, Meister M. 2003. Segregation of object and background motion in the retina. Nature 423: 401-408. Osten P, Khatri L, Perez JL, Kohr G, Giese G, et al. 2000. Mutagenesis reveals a role for ABP/GRIP binding to GluR2 in synaptic surface accumulation of the AMPA receptor. Neuron 27: 313-325. Osten P, Srivastava S, Inman GJ, Vilim FS, Khatri L, et al. 1998. The AMPA receptor GluR2 C terminus can mediate a reversible, ATP‐dependent interaction with NSF and alpha‐ and beta‐SNAPs. Neuron 21: 99-110. Oswald RE. 2004. Ionotropic glutamate receptor recognition and activation. Adv Protein Chem 68: 313-349. Park M, Penick EC, Edwards JG, Kauer JA, Ehlers MD. 2004. Recycling endosomes supply AMPA receptors for LTP. Science 305: 1972-1975. Partridge JG, Tang KC, Lovinger DM. 2000. Regional and postnatal heterogeneity of activity‐dependent long‐term changes in synaptic efficacy in the dorsal striatum. J Neurophysiol 84: 1422-1429. Passafaro M, Piech V, Sheng M. 2001. Subunit‐specific temporal and spatial patterns of AMPA receptor exocytosis in hippocampal neurons. Nat Neurosci 4: 917-926. Peineau S, Potier B, Petit F, Dournaud P, Epelbaum J, et al. 2003. AMPA‐sst2 somatostatin receptor interaction in rat hypothalamus requires activation of NMDA and/or metabotropic glutamate receptors and depends on intracellular calcium. J Physiol 546: 101-117. Pennartz CM, Dolleman‐Van der Weel MJ, Kitai ST, Lopes da Silva FH. 1992. Presynaptic dopamine D1 receptors attenuate excitatory and inhibitory limbic inputs to the shell region of the rat nucleus accumbens studied in vitro. J Neurophysiol 67: 1325-1334. Pennefather JN, Lecci A, Candenas ML, Patak E, Pinto FM, et al. 2004. Tachykinins and tachykinin receptors: A growing family. Life Sci 74: 1445-1463.

12

Perez JL, Khatri L, Chang C, Srivastava S, Osten P, et al. 2001. PICK1 targets activated protein kinase Calpha to AMPA receptor clusters in spines of hippocampal neurons and reduces surface levels of the AMPA‐type glutamate receptor subunit 2. J Neurosci 21: 5417-5428. Petralia RS, Zhao HM, Wang YX, Wenthold RJ. 1998. Variations in the tangential distribution of postsynaptic glutamate receptors in Purkinje cell parallel and climbing fiber synapses during development. Neuropharmacology 37: 1321-1334. Pinto A, Jankowski M, Sesack SR. 2003. Projections from the paraventricular nucleus of the thalamus to the rat prefrontal cortex and nucleus accumbens shell: Ultrastructural characteristics and spatial relationships with dopamine afferents. J Comp Neurol 459: 142-155. Plitzko D, Rumpel S, Gottmann K. 2001. Insulin promotes functional induction of silent synapses in differentiating rat neocortical neurons. Eur J Neurosci 14: 1412-1415. Pompeiano M, Palacios JM, Mengod G. 1994. Distribution of the serotonin 5‐HT2 receptor family mRNAs: Comparison between 5‐HT2A and 5‐HT2C receptors. Brain Res Mol Brain Res 23: 163-178. Popoli P, Pezzola A, Reggio R, Tiburzi F. 1999. Selective agonists of metabotropic glutamate receptors elicit significant EEG effects when infused in the nucleus accumbens of rats. Eur J Pharmacol 367: 183-188. Pow DV, Morris JF. 1989. Dendrites of hypothalamic magnocellular neurons release neurohypophysial peptides by exocytosis. Neuroscience 32: 435-439. Powers CJ, McLeskey SW, Wellstein A. 2000. Fibroblast growth factors, their receptors and signaling. Endocr Relat Cancer 7: 165-197. Price CJ, Kim P, Raymond LA. 1999. D1 dopamine receptor‐ induced cyclic AMP‐dependent protein kinase phosphorylation and potentiation of striatal glutamate receptors. J Neurochem 73: 2441-2446. Randic M, Cheng G, Kojic L. 1995. kappa‐opioid receptor agonists modulate excitatory transmission in substantia gelatinosa neurons of the rat spinal cord. J Neurosci 15: 6809-6826. Raymond JL, Lisberger SG, Mauk MD. 1996. The cerebellum: A neuronal learning machine. Science 272: 1126-1131. Raymond JR, Mukhin YV, Gelasco A, Turner J, Collinsworth G, et al. 2001. Multiplicity of mechanisms of serotonin receptor signal transduction. Pharmacol Ther 92: 179-212. Reynolds JN, Wickens JR. 2000. Substantia nigra dopamine regulates synaptic plasticity and membrane potential fluctuations in the rat neostriatum, in vivo. Neuroscience 99: 199-203. Risso F, Parodi M, Grilli M, Molfino F, Raiteri M, et al. 2004. Chronic nicotine causes functional upregulation of ionotropic glutamate receptors mediating hippocampal noradrenaline and striatal dopamine release. Neurochem Int 44: 293-301.

319

320

12

Regulation of AMPA receptors by metabotropic receptors and receptor tyrosine kinases

Roche KW, O’Brien RJ, Mammen AL, Bernhardt J, Huganir RL. 1996. Characterization of multiple phosphorylation sites on the AMPA receptor GluR1 subunit. Neuron 16: 1179-1188. Rodrigues Pdos S, Dowling JE. 1990. Dopamine induces neurite retraction in retinal horizontal cells via diacylglycerol and protein kinase C. Proc Natl Acad Sci USA 87: 9693-9697. Rogers TJ, Peterson PK. 2003. Opioid G protein‐coupled receptors: Signals at the crossroads of inflammation. Trends Immunol 24: 116-121. Rumbaugh G, Sia GM, Garner CC, Huganir RL. 2003. Synapse‐associated protein‐97 isoform‐specific regulation of surface AMPA receptors and synaptic function in cultured neurons. J Neurosci 23: 4567-4576. Rusin KI, Ryu PD, Randic M. 1992. Modulation of excitatory amino acid responses in rat dorsal horn neurons by tachykinins. J Neurophysiol 68: 265-286. Rusin KI, Jiang MC, Cerne R, Randic M. 1993. Interactions between excitatory amino acids and tachykinins in the rat spinal dorsal horn. Brain Res Bull 30: 329-338. Rutherford LC, Nelson SB, Turrigiano GG. 1998. BDNF has opposite effects on the quantal amplitude of pyramidal neuron and interneuron excitatory synapses. Neuron 21: 521-530. Saal D, Dong Y, Bonci A, Malenka RC. 2003. Drugs of abuse and stress trigger a common synaptic adaptation in dopamine neurons. Neuron 37: 577-582. Salt TE. 1987. Excitatory amino acid receptors and synaptic transmission in the rat ventrobasal thalamus. J Physiol 391: 499-510. Saltiel AR, Pessin JE. 2002. Insulin signaling pathways in time and space. Trends Cell Biol 12: 65-71. Sandkuhler J. 1996. The organization and function of endogenous antinociceptive systems. Prog Neurobiol 50: 49-81. Schmidt KF, Kruse M, Hatt H. 1994. Dopamine alters glutamate receptor desensitization in retinal horizontal cells of the perch (Perca fluviatilis). Proc Natl Acad Sci USA 91: 8288-8291. Schnell E, Sizemore M, Karimzadegan S, Chen L, Bredt DS, et al. 2002. Direct interactions between PSD‐95 and stargazin control synaptic AMPA receptor number. Proc Natl Acad Sci USA 99: 13902-13907. Schoepp DD, Jane DE, Monn JA. 1999. Pharmacological agents acting at subtypes of metabotropic glutamate receptors. Neuropharmacology 38: 1431-1476. Seeburg PH. 1996. The role of RNA editing in controlling glutamate receptor channel properties. J Neurochem 66: 1-5. Seeburg PH, Hartner J. 2003. Regulation of ion channel/ neurotransmitter receptor function by RNA editing. Curr Opin Neurobiol 13: 279-283.

Seeburg PH, Higuchi M, Sprengel R. 1998. RNA editing of brain glutamate receptor channels: Mechanism and physiology. Brain Res Brain Res Rev 26: 217-229. Sesack SR, Pickel VM. 1990. In the rat medial nucleus accumbens, hippocampal and catecholaminergic terminals converge on spiny neurons and are in apposition to each other. Brain Res 527: 266-279. Sesack SR, Pickel VM. 1992. Prefrontal cortical efferents in the rat synapse on unlabeled neuronal targets of catecholamine terminals in the nucleus accumbens septi and on dopamine neurons in the ventral tegmental area. J Comp Neurol 320: 145-160. Severini C, Improta G, Falconieri‐Erspamer G, Salvadori S, Erspamer V. 2002. The tachykinin peptide family. Pharmacol Rev 54: 285-322. Shen L, Liang F, Walensky LD, Huganir RL. 2000. Regulation of AMPA receptor GluR1 subunit surface expression by a 4.1N‐linked actin cytoskeletal association. J Neurosci 20: 7932-7940. Sherrard RM, Richardson NA, Sara VR. 1997. Localisation of insulin‐like growth factor‐I (IGF‐I) immunoreactivity in the olivocerebellar system of developing and adult rats. Brain Res Dev Brain Res 98: 102-113. Sherwood NT, Lo DC. 1999. Long‐term enhancement of central synaptic transmission by chronic brain‐derived neurotrophic factor treatment. J Neurosci 19: 7025-7036. Shi S, Hayashi Y, Esteban JA, Malinow R. 2001. Subunit‐ specific rules governing AMPA receptor trafficking to synapses in hippocampal pyramidal neurons. Cell 105: 331-343. Shibuki K, Kimura S. 1997. Dynamic properties of nitric oxide release from parallel fibres in rat cerebellar slices. J Physiol 498 (Pt 2): 443-452. Shigemoto R, Abe T, Nomura S, Nakanishi S, Hirano T. 1994. Antibodies inactivating mGluR1 metabotropic glutamate receptor block long‐term depression in cultured Purkinje cells. Neuron 12: 1245-1255. Shutoh F, Hamada S, Shibata M, Narita M, Shiga T, et al. 2000. Long term depletion of serotonin leads to selective changes in glutamate receptor subunits. Neurosci Res 38: 365-371. Smith Y, Bennett BD, Bolam JP, Parent A, Sadikot AF. 1994. Synaptic relationships between dopaminergic afferents and cortical or thalamic input in the sensorimotor territory of the striatum in monkey. J Comp Neurol 344: 1-19. Snyder GL, Allen PB, Fienberg AA, Valle CG, Huganir RL, et al. 2000. Regulation of phosphorylation of the GluR1 AMPA receptor in the neostriatum by dopamine and psychostimulants in vivo. J Neurosci 20: 4480-4488. Snyder EM, Philpot BD, Huber KM, Dong X, Fallon JR, et al. 2001. Internalization of ionotropic glutamate receptors in response to mGluR activation. Nat Neurosci 4: 1079-1085.

Regulation of AMPA receptors by metabotropic receptors and receptor tyrosine kinases Sobolevsky AI, Yelshansky MV, Wollmuth LP. 2003. Different gating mechanisms in glutamate receptor and Kþchannels. J Neurosci 23: 7559-7568. Soderling TR, Derkach VA. 2000. Postsynaptic protein phosphorylation and LTP. Trends Neurosci 23: 75-80. Sommer B, Kohler M, Sprengel R, Seeburg PH. 1991. RNA editing in brain controls a determinant of ion flow in glutamate‐gated channels. Cell 67: 11-19. Sommer B, Keinanen K, Verdoorn TA, Wisden W, Burnashev N, et al. 1990. Flip and flop: A cell‐specific functional switch in glutamate‐operated channels of the CNS. Science 249: 1580-1585. Song I, Huganir RL. 2002. Regulation of AMPA receptors during synaptic plasticity. Trends Neurosci 25: 578-588. Song I, Kamboj S, Xia J, Dong H, Liao D, et al. 1998. Interaction of the N‐ethylmaleimide‐sensitive factor with AMPA receptors. Neuron 21: 393-400. Srivastava S, Osten P, Vilim FS, Khatri L, Inman G, et al. 1998. Novel anchorage of GluR2/3 to the postsynaptic density by the AMPA receptor‐binding protein ABP. Neuron 21: 581-591. Standaert DG, Needleman P, Saper CB. 1986. Organization of atriopeptin‐like immunoreactive neurons in the central nervous system of the rat. J Comp Neurol 253: 315-341. Steinbusch HW. 1981. Distribution of serotonin‐immunoreactivity in the central nervous system of the rat‐cell bodies and terminals. Neuroscience 6: 557-618. Stern JE, Galarreta M, Foehring RC, Hestrin S, Armstrong WE. 1999. Differences in the properties of ionotropic glutamate synaptic currents in oxytocin and vasopressin neuroendocrine neurons. J Neurosci 19: 3367-3375. Sugiura Y, Lee CL, Perl ER. 1986. Central projections of identified, unmyelinated (C) afferent fibers innervating mammalian skin. Science 234: 358-361. Sugiura Y, Terui N, Hosoya Y. 1989. Difference in distribution of central terminals between visceral and somatic unmyelinated (C) primary afferent fibers. J Neurophysiol 62: 834-840. Svenningsson P, Tzavara ET, Witkin JM, Fienberg AA, Nomikos GG, et al. 2002. Involvement of striatal and extrastriatal DARPP‐32 in biochemical and behavioral effects of fluoxetine (Prozac). Proc Natl Acad Sci USA 99: 31823187. Swanson CJ, Kalivas PW. 2000. Regulation of locomotor activity by metabotropic glutamate receptors in the nucleus accumbens and ventral tegmental area. J Pharmacol Exp Ther 292: 406-414. Swayze RD, Lise MF, Levinson JN, Phillips A, El‐Husseini A. 2004. Modulation of dopamine mediated phosphorylation of AMPA receptors by PSD‐95 and AKAP79/150. Neuropharmacology 47: 764-778.

12

Taverna S, Pennartz CM. 2003. Postsynaptic modulation of AMPA‐ and NMDA‐receptor currents by Group III metabotropic glutamate receptors in rat nucleus accumbens. Brain Res 976: 60-68. Testa CM, Standaert DG, Young AB, Penney JB Jr. 1994. Metabotropic glutamate receptor mRNA expression in the basal ganglia of the rat. J Neurosci 14: 3005-3018. Thomas MJ, Beurrier C, Bonci A, Malenka RC. 2001. Long‐ term depression in the nucleus accumbens: A neural correlate of behavioral sensitization to cocaine. Nat Neurosci 4: 1217-1223. Thomas MJ, Moody TD, Makhinson M, O’Dell TJ. 1996. Activity‐dependent beta‐adrenergic modulation of low frequency stimulation induced LTP in the hippocampal CA1 region. Neuron 17: 475-482. Timney B, Mitchell DE, Giffin F. 1978. The development of vision in cats after extended periods of dark‐rearing. Exp Brain Res 31: 547-560. Tomita S, Fukata M, Nicoll RA, Bredt DS. 2004. Dynamic interaction of stargazin‐like TARPs with cycling AMPA receptors at synapses. Science 303: 1508-1511. Tomita S, Chen L, Kawasaki Y, Petralia RS, Wenthold RJ, et al. 2003. Functional studies and distribution define a family of transmembrane AMPA receptor regulatory proteins. J Cell Biol 161: 805-816. Totterdell S, Smith AD. 1989. Convergence of hippocampal and dopaminergic input onto identified neurons in the nucleus accumbens of the rat. J Chem Neuroanat 2: 285-298. Trudeau LE, Castellucci VF. 1995. Postsynaptic modifications in long‐term facilitation in Aplysia: Upregulation of excitatory amino acid receptors. J Neurosci 15: 1275-1284. Trueblood PR, Levine MS, Chandler SH. 1996. Dual‐component excitatory amino acid‐mediated responses in trigeminal motoneurons and their modulation by serotonin in vitro. J Neurophysiol 76: 2461-2473. Tseng KY, O’Donnell P. 2004. Dopamine‐glutamate interactions controlling prefrontal cortical pyramidal cell excitability involve multiple signaling mechanisms. J Neurosci 24:5131-5139 [In this study, the modulatory actions of D1‐ and D2‐receptors on prefrontal cortex pyramidal cell excitability were assessed using whole‐cell patch‐clamp recordings in brain slices obtained from developmentally mature rats. D2‐receptor activation was found to attenuate the excitatory effects of AMPA, through a process involving postsynaptic inhibition of PKA and activation of phospholipase C.]. Uchimura N, Higashi H, Nishi S. 1989. Membrane properties and synaptic responses of the guinea pig nucleus accumbens neurons in vitro. J Neurophysiol 61: 769-779. Uemura T, Mori H, Mishina M. 2004. Direct interaction of GluRdelta2 with Shank scaffold proteins in cerebellar Purkinje cells. Mol Cell Neurosci 26: 330-341.

321

322

12

Regulation of AMPA receptors by metabotropic receptors and receptor tyrosine kinases

Ugolini A, Corsi M, Bordi F. 1997. Potentiation of NMDA and AMPA responses by group I mGluR in spinal cord motoneurons. Neuropharmacology 36: 1047-1055. Ugolini A, Corsi M, Bordi F. 1999. Potentiation of NMDA and AMPA responses by the specific mGluR5 agonist CHPG in spinal cord motoneurons. Neuropharmacology 38: 1569-1576. Umemiya M, Raymond LA. 1997. Dopaminergic modulation of excitatory postsynaptic currents in rat neostriatal neurons. J Neurophysiol 78: 1248-1255. Undie AS, Friedman E. 1990. Stimulation of a dopamine D1 receptor enhances inositol phosphates formation in rat brain. J Pharmacol Exp Ther 253: 987-992. Undie AS, Friedman E. 1992. Selective dopaminergic mechanism of dopamine and SKF38393 stimulation of inositol phosphate formation in rat brain. Eur J Pharmacol 226: 297-302. Undie AS, Weinstock J, Sarau HM, Friedman E. 1994. Evidence for a distinct D1‐like dopamine receptor that couples to activation of phosphoinositide metabolism in brain. J Neurochem 62: 2045-2048. Ungless MA, Whistler JL, Malenka RC, Bonci A. 2001. Single cocaine exposure in vivo induces long‐term potentiation in dopamine neurons. Nature 411: 583-587. Vanhoose AM, Winder DG. 2003. NMDA and beta1‐adrenergic receptors differentially signal phosphorylation of glutamate receptor type 1 in area CA1 of hippocampus. J Neurosci 23: 5827-5834. Versaux‐Botteri C, Gibert JM, Nguyen‐Legros J, Vernier P. 1997. Molecular identification of a dopamine D1b receptor in bovine retinal pigment epithelium. Neurosci Lett 237: 9-12. Veruki ML. 1997. Dopaminergic neurons in the rat retina express dopamine D2/3 receptors. Eur J Neurosci 9: 1096-1100. Veruki ML, Wassle H. 1996. Immunohistochemical localization of dopamine D1 receptors in rat retina. Eur J Neurosci 8: 2286-2297. Viegi A, Cotrufo T, Berardi N, Mascia L, Maffei L. 2002. Effects of dark rearing on phosphorylation of neurotrophin Trk receptors. Eur J Neurosci 16: 1925-1930. Walaas SI, Aswad DW, Greengard P. 1983. A dopamine‐ and cyclic AMP‐regulated phosphoprotein enriched in dopamine‐innervated brain regions. Nature 301: 69-71. Wang F, McKeehan WL. 2003. The Fibroblast Growth Factor (FGF) Signaling Complex. Handbook of Cell Signaling ‐ CD‐ROM, Vol. 1. Bradshaw RA, Dennis EA, editors. Elsevier, Academic Press. Wang GD, Zhuo M. 2002. Synergistic enhancement of glutamate‐mediated responses by serotonin and forskolin in adult mouse spinal dorsal horn neurons. J Neurophysiol 87: 732-739. Wang LY, Dudek EM, Browning MD, Mac Donald JF. 1994. Modulation of AMPA/kainate receptors in cultured murine

hippocampal neurones by protein kinase C. J Physiol 475: 431-437. Wang XF, Daw NW. 1996. Metabotropic glutamate receptors potentiate responses to NMDA and AMPA from layer V cells in rat visual cortex. J Neurophysiol 76: 808-815. Wang XF, Daw NW, Jin X. 1998. The effect of ACPD on the responses to NMDA and AMPA varies with layer in slices of rat visual cortex. Brain Res 812: 186-192. Wang YT, Linden DJ. 2000. Expression of cerebellar long‐term depression requires postsynaptic clathrin‐mediated endocytosis. Neuron 25: 635-647 [This paper shows that interference with the clathrin endocytic complex, by the postsynaptic application of dynamin and amphiphysin peptides, blocks LTD expression in cultured Purkinje neurons.]. Werther GA, Abate M, Hogg A, Cheesman H, Oldfield B, et al. 1990. Localization of insulin‐like growth factor‐I mRNA in rat brain by in situ hybridization–relationship to IGF‐I receptors. Mol Endocrinol 4: 773-778. Wickelgren I. 1998. Tracking insulin to the mind. Science 280: 517-519. Winder DG, Martin KC, Muzzio IA, Rohrer D, Chruscinski A, et al. 1999. ERK plays a regulatory role in induction of LTP by theta frequency stimulation and its modulation by beta‐ adrenergic receptors. Neuron 24: 715-726. Witkovsky P. 2004. Dopamine and retinal function. Doc Ophthalmol 108: 17-40. Witkovsky P, Stone S, Besharse JC. 1988. Dopamine modifies the balance of rod and cone inputs to horizontal cells of the Xenopus retina. Brain Res 449: 332-336. Wolf ME. 1998. The role of excitatory amino acids in behavioral sensitization to psychomotor stimulants. Prog Neurobiol 54: 679-720. Wolf ME, Mangiavacchi S, Sun X. 2003. Mechanisms by which dopamine receptors may influence synaptic plasticity. Ann NY Acad Sci 1003: 241-249. Womack MD, Mac Dermott AB, Jessell TM. 1988. Sensory transmitters regulate intracellular calcium in dorsal horn neurons. Nature 334: 351-353. Wozniak M, Rydzewski B, Baker SP, Raizada MK. 1993. The cellular and physiological actions of insulin in the central nervous system. Neurochem Int 22: 1-10. Xia J, Zhang X, Staudinger J, Huganir RL. 1999. Clustering of AMPA receptors by the synaptic PDZ domain‐containing protein PICK1. Neuron 22: 179-187. Xia J, Chung HJ, Wihler C, Huganir RL, Linden DJ. 2000. Cerebellar long‐term depression requires PKC‐regulated interactions between GluR2/3 and PDZ domain‐containing proteins. Neuron 28: 499-510 [This paper shows that perfusion of cultured cerebellar Purkinje neurons with peptides corresponding to the phosphorylated and dephosphorylated GluR2 C‐terminal PDZ‐binding motif,

Regulation of AMPA receptors by metabotropic receptors and receptor tyrosine kinases which block the interaction of GluR2 with PICK1 or with GRIP and PICK1, respectively, attenuates LTD induction.]. Xiao MY, Zhou Q, Nicoll RA. 2001. Metabotropic glutamate receptor activation causes a rapid redistribution of AMPA receptors. Neuropharmacology 41: 664-671. Yan Z, Hsieh‐Wilson L, Feng J, Tomizawa K, Allen PB, et al. 1999. Protein phosphatase 1 modulation of neostriatal AMPA channels: Regulation by DARPP‐32 and spinophilin. Nat Neurosci 2: 13-17 [Using neostriatal neurons in culture, the authors of this study demonstrate that the rundown of AMPAR‐mediated currents is prevented by the activation of D1‐receptors, or the inhibition of phosphatase 1/2A. The link between the two observations is provided by the fact that DARR‐32, a protein enriched in the neostritum, which in its phosphorylated form is an inhibitor of protein phosphatase‐1, is phosphorylated following D1‐receptor activation.]. Yang CR, Mogenson GJ. 1984. Electrophysiological responses of neurones in the nucleus accumbens to hippocampal stimulation and the attenuation of the excitatory responses by the mesolimbic dopaminergic system. Brain Res 324: 69-84. Yim CY, Mogenson GJ. 1982. Response of nucleus accumbens neurons to amygdala stimulation and its modification by dopamine. Brain Res 239: 401-415. Yoshimura M, Jessell TM. 1989. Primary afferent‐evoked synaptic responses and slow potential generation in rat substantia gelatinosa neurons in vitro. J Neurophysiol 62: 96-108.

12

Yoshimura M, Jessell T. 1990. Amino acid‐mediated EPSPs at primary afferent synapses with substantia gelatinosa neurones in the rat spinal cord. J Physiol 430: 315-335. Yu J, Ustach C, Kim HR. 2003. Platelet‐derived growth factor signaling and human cancer. J Biochem Mol Biol 36: 49-59. Yuzaki M. 2004. The delta2 glutamate receptor: A key molecule controlling synaptic plasticity and structure in Purkinje cells. Cerebellum 3: 89-93. Zho WM, You JL, Huang CC, Hsu KS. 2002. The group I metabotropic glutamate receptor agonist (S)‐3,5‐dihydroxyphenylglycine induces a novel form of depotentiation in the CA1 region of the hippocampus. J Neurosci 22: 8838-8849. Zhou Q, Xiao M, Nicoll RA. 2001. Contribution of cytoskeleton to the internalization of AMPA receptors. Proc Natl Acad Sci USA 98: 1261-1266. Zhu JJ, Esteban JA, Hayashi Y, Malinow R. 2000. Postnatal synaptic potentiation: Delivery of GluR4‐containing AMPA receptors by spontaneous activity. Nat Neurosci 3: 10981106. Zhuo M, Gebhart GF. 1992. Characterization of descending facilitation and inhibition of spinal nociceptive transmission from the nuclei reticularis gigantocellularis and gigantocellularis pars alpha in the rat. J Neurophysiol 67: 1599-1614. Zhuo M, Gebhart GF. 1997. Biphasic modulation of spinal nociceptive transmission from the medullary raphe nuclei in the rat. J Neurophysiol 78: 746-758.

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Taurine in Neurotransmission

P. Saransaari . S. S. Oja

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Introduction and Neurotransmitter Criteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 326

2 2.1 2.2 2.3

Criterion of Presence . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 326 Occurrence and Distribution . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 326 Biosynthesis and Catabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 327 Uptake . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 328

3 Physiological and Pharmacological Actions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 328 3.1 Effects on Membrane Ion Conductances . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 328 3.2 Putative Taurine Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 329 4

Taurine Release . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 330

5 5.1 5.2 5.3 5.4

Neuromodulatory Functions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 331 Interactions with GABAergic Systems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 331 Interactions with Glycinergic Systems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 332 Interactions with Other Transmitter Systems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 333 Effects on Calcium Levels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 334

6

Conclusions and Perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 334

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2008 Springer ScienceþBusiness Media, LLC.

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Abstract: This chapter reviews present knowledge of the possible roles of taurine as a neurotransmitter or neuromodulator. Neurons and glia possess biosynthetic machinery and ample amounts and efficient reuptake of taurine. It is released by depolarization, but the Ca2þ dependency of stimulated release is as yet not definitely settled. Taurine enhances the chloride conductance of plasma membranes in nerve cells and induces hyperpolarization with subsequent inhibition. It remains open whether or not taurine possesses receptors of its own or whether its actions are mediated by GABA and glycine receptors. Taurine may be a neurotransmitter in certain brain areas, more likely in developing animals and in species other than mammals, but no taurinergic nerve tracts are known. Taurine is not a neuromodulator in the classical sense as it has actions of its own in the synaptic region and does not of itself influence the functions of established neurotransmitters. List of Abbreviations: CNS, central nervous system; GABA, g‐aminobutyrate; MPPþ, 1‐methyl‐4‐phenylpyridinium; TAG, 6‐aminomethyl‐3‐methyl‐4H‐1,2,4‐benzothiadiazine‐1,1‐dioxide hydrogen maleate; TAUTs, taurine transporters

1

Introduction and Neurotransmitter Criteria

Since the 1960s, the sulfur‐containing amino acid taurine (2‐aminoethanesulfonic acid) has been assigned a number of important functions in the central nervous system (CNS), one of the first being a role as an inhibitory neurotransmitter (see Oja and La¨hdesma¨ki, 1974; Oja et al., 1977, and references therein). It has since been envisaged as a membrane stabilizer, endogenous antiepileptic agent, cell volume regulator, and neuromodulator and neuroprotector, and thought to maintain calcium homeostasis and membrane integrity. We have therefore authored an exhaustive review on these putative actions of taurine in this Handbook edition (Volume 6: Amino Acids and Peptides in the Nervous System). Here, we focus on the role of taurine in neurotransmission and endeavor to draw inferences as to whether it could qualify as a neurotransmitter or neuromodulator. A neurotransmitter should fulfill four criteria according to one traditional classification: (1) presence, (2) physiological identity, (3) pharmacological identity, and (4) chemical identity (Steiner, 1971). Accordingly: (1) A transmitter must be present in the synaptic region with enzymes with its biosynthetic machinery preferably nearby. Mechanisms for its inactivation must exist, fast catabolism or efficient re‐uptake. (2) The transmitter should at physiological concentrations alter the polarization and ion conductances of the postsynaptic membrane. Isolation and characterization of postsynaptic receptors are important evidences. (3) Drugs should similarly affect physiological impulse transmission and the postsynaptic responses evoked by the transmitter candidate. (4) The transmitter should be released from nerve terminals and extracellularly recovered and identified upon physiological impulse transmission. On the other hand, a neuromodulator has no function of its own in synaptic transmission, but it affects synaptic events otherwise, interfering, for example, with the actions of proper transmitters.

2

Criterion of Presence

2.1 Occurrence and Distribution Taurine is a free amino acid abundant in the CNS, only glutamate being generally present at higher concentrations (Oja and Kontro, 1983; Huxtable, 1989). There are marked species differences, the regional distribution is somewhat heterogeneous, and a gradual decrease in taurine occurs during postnatal development in most animal species (Oja et al., 1977; Oja and Kontro, 1983; Huxtable, 1989, and references in them). The magnitude of this postnatal decrease varies from area to area, being several times greater in the spinal cord and medulla than, for example, in the cerebellar and cerebral cortex (Cutler and Dudzinski, 1974). The cerebral cortex, cerebellum, olfactory bulbs, striatum, and hypothalamus contain more taurine than the pons–medulla and spinal cord. In individual nuclei, taurine is enriched in the lateral geniculate

Taurine in neurotransmission

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and inferior colliculus. In the latter nucleus, the posterior region has the highest and the anterior region the lowest taurine concentration in the cat (Guidotti et al., 1972). Taurine distribution in the rat spinal cord and thalamus is fairly even, whereas the canine spinal cord shows segmental differences (Lane et al., 1978). The intraregional distribution of taurine in the rat cerebellum was thoroughly analyzed at a fairly early date. There was more taurine in the inhibitory stellate cells, which were at the time assumed to use it as a transmitter (McBride and Frederickson, 1980), than in other cerebellar cortical cells (Nadi et al., 1977). In the bovine brain, taurine is enriched in synaptic vesicles (Kontro et al., 1980), but it remains open whether it resides in the vesicular fluid, since taurine has an inherent tendency to bind relatively tightly to synaptic membranes (Marnela et al., 1980; Kontro and Oja, 1987a; Frosini et al., 2003). The introduction of taurine‐specific antibodies and subsequent methodological developments have enabled extensive mapping of taurine‐containing cells and cell organelles in the CNS. In principle, all cell types have been reported to contain taurine (Huxtable, 1989). However, in the cat perihypoglossal and vestibular nuclei only glial cells immunostained for taurine, but Purkinje cell axons also exhibited a positive reaction (Yingcharoen et al., 1989; Walberg et al., 1990). In the rat, taurine‐like immunoreactivity has also been observed in Purkinje cell bodies and their dendrites, mossy fibers, and Golgi axons (Nagelhuis et al., 1993; Gragera et al., 1995), but glial cells and their processes also exhibit positive staining (Torp et al., 1991). In the rat vestibular nuclei, the taurine and g‐aminobutyrate (GABA) distributions show a significant correlation (Li et al., 1994), and unilateral vestibular ganglionectomy affects them both similarly (Li et al., 1996). No such correlation has been seen in the rat auditory, olfactory, and visual systems (Ross et al., 1995). The taurine concentration diminishes from superficial to deep layers in the dorsal cochlear nucleus (Godfrey et al., 2000), and in the olfactory bulb taurine is enriched in primary olfactory neurons, taurine‐like immunoreactivity being more intense in their axons than in the terminals (Didier et al., 1994). In the supraoptic nucleus, immunoreactivity has been found to be intense in glial cell bodies in the ventral glial lamina and in the glial processes surrounding magnocellular neurons (Decavel and Hatton, 1995).

2.2 Biosynthesis and Catabolism The brain contains the enzymatic machinery for taurine biosynthesis from its precursor amino acids. The sulfur‐containing amino acids methionine and cysteine serve as precursors. In two major synthesis pathways, cysteine is either first oxidized by cysteine dioxygenase (EC 1.13.11.20) to 3‐sulfinoalanine (cysteinesulfinate), which is then decarboxylated by sulfino decarboxylase (EC 4.1.1.29, also known as cysteinesulfinate decarboxylase) to yield hypotaurine (Huxtable, 1986), or transformed by a more devious route via coenzyme A to pantetheine and then to cysteamine, which is converted to hypotaurine by means of cysteamine dioxygenase (EC 1.13.11.19) (Coloso et al., 2006). The preferred route is probably cysteine decarboxylation. The final step, oxidation of hypotaurine to taurine, may be nonenzymatic oxidation (Fellman and Roth, 1985), though enzyme(s) can also be involved (Oja and Kontro, 1981; Kontro and Oja, 1985a). The above biosynthetic routes and the enzymes involved have been thoroughly described in earlier reviews by Oja and Kontro (1983), Huxtable (1989), and Tappaz (2004). However, taurine biosynthesis in situ may be slower than supply from other organs (Beetsch and Olson, 1996, 1998), particularly from the liver (Stipanuk et al., 2002; Tappaz, 2004). In our opinion the biosynthetic machinery in the brain supplies enough taurine for synaptic functions. However, some species evince an inadequate capability to synthesize taurine to meet the body demands. For such animals, e.g., carnivorous cats, taurine is an essential nutrient and its deficiency results in grave structural malformations and functional deficits (Sturman et al., 1985; Sturman, 1993; Imaki et al., 1986; Maar et al., 1995). Also, in primates and humans taurine is an essential nutrient for babies (Neuringer et al., 1985; Imaki et al., 1987), its deficiency resulting in similar developmental problems (Geggel et al., 1985; Ament et al., 1986). Taurine is not broken down in mammalian organisms. It is conjugated with bile acids in the liver, and the kidneys excrete taurine into the urine, the rate of excretion reflecting the dietary intake and plasma levels (Chesney et al., 1985). Taurine supplementation increases brain taurine only moderately (Satsu et al.,

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2002; Lallemand and De Witte, 2004), and only temporarily (Salima¨ki et al., 2003). The slow breakdown and elimination of taurine signifies that these cannot play any role in the termination of its possible synaptic action in the CNS.

2.3 Uptake The properties of cellular taurine have been extensively described and discussed in reviews by Oja and Kontro (1983) and Huxtable (1989). The uptake is saturable, concentrative, energy‐, Naþ‐, and Cl
‐dependent, and structurally specific for o‐amino acids (or b‐amino acids), i.e., the acidic group and basic groups should reside at the opposite ends of the molecule, separated by two or three carbon atoms. Two or three Naþ ions are required for the transport of one taurine molecule. Both neurons and glial cells actively take up taurine from the extracellular spaces. The uptake comprises two components, low and high affinity (Kontro and Oja, 1978; Hanretta and Lombardini, 1987). The estimated Km constants for the low‐affinity transport have been within the micromolar range and those for high‐affinity transport within the millimolar range. These constants are generally smaller in preparations from the developing than the adult brain. Taurine transporters (TAUTs) have been cloned from different CNS preparations (Liu et al., 1992; Smith et al., 1992; Ramamoorthy et al., 1994). They show marked homology with each other and also with the transporters of established neurotransmitters (Smith et al., 1992; Uchida et al., 1992). The distribution of mRNA for the rat TAUT is more or less universal within the CNS and among different types of cells, both glial cells and neurons (Borden et al., 1995; Lake and Orlowski, 1996; Kang, 2000; Olson and Martinho, 2006), though the two different clones TAUT‐1 and TAUT‐2 are differently localized (Pow et al., 2002). TAUT‐1 is detectable in the rat pituicytes, cerebellar Purkinje cells, and in photoreceptors and bipolar cells in the retina. TAUT‐2 is more widely distributed in the CNS, being however predominantly associated with Bergmann glial cells in the cerebellum and with astrocytes in other brain areas and, at lower levels, with some neurons such as CA1 pyramidal cells in the hippocampus (Pow et al., 2002). In the mouse cerebral cortex, TAUT has been shown to be responsible for uptake, its expression being increased with development (Fujita et al., 2006). Electrophysiological experiments have also confirmed the presence of TAUTs in Bergmann glia (Barakat et al., 2002). TAUT exhibits the same electrogenic stoichiometry as the GABA transporter GAT1 (Beckman and Quick, 1998). Its gene expression appears to be under the control of two antagonistic regulations, osmolarity‐induced upregulation and taurine‐induced downregulation (Bitoun and Tappaz, 2000), but the regulation in the brain is still a partially open matter. The most potent inhibitors of cloned TAUTs are hypotaurine and b‐alanine, (Kulanthaivel et al., 1991; Liu et al., 1992; Smith et al., 1992; Vinnakota et al., 1997), which also most efficiently inhibit taurine transport in brain slices (La¨hdesma¨ki and Oja, 1973) and cultured cells (Holopainen et al., 1983; Rebel et al., 1994). The transporters also accept GABA, albeit with a much lesser affinity (Vinnakota et al., 1997). On the other hand, the cloned GABA transporters also transport taurine, their affinity for taurine being however much lower than that for GABA (Liu et al., 1993; Melamed and Kanner, 2004).

3

Physiological and Pharmacological Actions

3.1 Effects on Membrane Ion Conductances Taurine has been shown to inhibit neuronal firing by producing hyperpolarization via alteration of membrane permeability to ions in different brain areas, including the cerebral cortex, diencephalon, striatum, hippocampus, cerebellum, brain stem, and spinal cord (see references in Oja and Kontro, 1978, 1983; Huxtable, 1989). The actions of taurine are in most cases blocked by antagonists of GABA and/or glycine receptors, depending on the brain area. In early experiments, taurine effects were generally abolished by GABA antagonists in the higher brain areas and by glycine antagonists in the brain stem and spinal cord. Since there are no sufficiently specific taurine antagonists available, except for 6‐aminomethyl‐3‐methyl‐ 4H‐1,2,4‐benzothiadiazine‐1,1‐dioxide hydrogen maleate (TAG) (Yarborough et al., 1981), the effects of

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taurine per se are difficult to demarcate from those mediated via GABA or glycine receptors (see also Chapters 3.2, 5.1, and 5.2). However, it has at least been shown that the taurine antagonist TAG blocks taurine‐induced hyperpolarization in Purkinje cell dendrites in the guinea pig cerebellum (Okamoto et al., 1983b, c). The rigid analogs of taurine, 3‐amino‐1,2,4-benzothiadiazine-1,1-dioxide and 3‐aminomethyl‐ 1,2,4‐benzothiadiazine‐1,1‐dioxide, have been claimed to possess better selectivity than TAG (Braghiroli et al., 1990), but to our best knowledge they have not since been applied in other investigations. We have shown that at low concentrations taurine inhibits GABA‐induced Cl
fluxes into synaptic membrane microsacs prepared from different brain areas, but at higher concentrations enhances the fluxes, apparently due to activatory properties of its own (Oja et al., 1990b). In keeping with such observations, taurine increases Cl
conductance in cerebellar Purkinje cell dendrites in vitro (Okamoto et al., 1983a), in rat substantia nigra pars reticulate neurons (Ye et al., 1997), and in mitral and tufted cells in slices from the rat main olfactory bulb (Puopolo et al., 1998). In these cells, the currents evoked by GABA are fast and the currents induced by taurine slow (Belluzzi et al., 2004). Taurine may thus cause prolonged reduction in the input resistance and sustain a prolonged and modest inhibition of these cells. It has also induced single‐ channel currents by opening Cl
channels in cultured rat cerebellar granule cells in patch‐clamp experiments (Linne et al., 1996). In dopamine neurons in the rat substantia nigra (Ha¨user et al., 1992), freshly isolated neurons from the medulla oblongata and hippocampus (Krishtal et al., 1988), the rat sacral dorsal commissural neurons (Wang et al., 1998), and in acutely dissociated rat hippocampal neurons (Yu et al., 2003) taurine and glycine activate the same Cl
conductance and apparently act at least predominantly at the same recognition site. Together with b‐alanine, taurine also activates glycine receptors in cultured rat hippocampal slices (Mori et al., 2002). Isolated rat basolateral amygdala neurons respond to extracellular application of taurine with pronounced changes in the resting membrane current (McCool and Botting, 2000). Taurine increases the amplitude of spontaneous inhibitory postsynaptic currents in dopaminergic neurons in the ventral tegmental area in young rats, in which GABA is excitatory due to variations in the extracellular content of Cl
, with a potency of one‐ tenth that of glycine (Ye et al., 2004). By reason of developmental changes in the extracellular concentration of Cl
, the activation of Cl
channels by taurine (Yoshida et al., 2004) during cortical circuit formation in the postnatally developing mouse alters its physiological effects from excitatory to inhibitory. Within the same time frame the target of taurine shifts from glycine receptors to GABA receptors (Yoshida et al., 2004).

3.2 Putative Taurine Receptors The possible neurotransmitter function of taurine presupposes the existence of membrane receptors. As stated above, it has been difficult to determine whether the physiological functions of taurine are mediated by GABA and/or glycine receptors or whether specific taurine receptors are involved in certain brain areas, at particular developmental stages and in different animal species. Taurine binding to brain membranes has likewise proved difficult to demonstrate. We were the first to report taurine binding to mouse brain synaptic membranes (Kontro and Oja, 1983), if the membranes were subjected to freezing–thawing cycles and detergent treatments to extract as much as possible of the endogenous taurine attached to the preparations tested (Kontro and Oja, 1987a). The binding proved to be Naþ independent and exhibited sigmoidal dependence on the ligand concentration (Kontro and Oja, 1985b, 1987d). The taurine antagonist TAG proved to be an effective displacer of this binding, but it was also affected by the glycine receptor antagonist strychnine and by GABA antagonists. It was therefore not possible to demarcate the binding to glycine and GABA receptors from the possible binding to taurine receptors. In the frog spinal cord, two taurine receptor subtypes were surmised to exist based on their pharmacological properties, differing from the responses to GABA and glycine (Kudo et al., 1988). In the olfactory organ of the lobster, preliminary identification and partial characterization of the putative taurine receptor proteins have been made (Sung et al., 1996). Frosini and coworkers (2003) present a very thorough characterization of [3H]taurine binding to washed and detergent‐treated synaptic membranes from the rabbit brain. They showed that taurine binding was displaced by TAG, 2‐aminoethylarsonate, 2‐hydroxyethanesulfonate, and ()cis‐2‐aminocyclohexanesulfonate. These effectors did not interact with GABAA

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and GABAB receptors, taurine and GABA uptake systems, or GABA transaminase. The taurine binding studied does not thus represent binding to GABA receptors, but the effects of glycine agonists and antagonists on this taurine binding were not tested. As we (Kontro and Oja, 1987a) and Frosini and his coworkers (2003) have stressed, detergents are a must in removing endogenous binding inhibitors such as taurine itself from synaptic membranes, as was the case with early studies on GABAA receptors (Napias et al., 1980). Only using detergents can specific binding of taurine be demonstrated. Moreover, the endogenous muscimol‐binding inhibitor in brain tissue has been claimed to a more potent inhibitor of taurine binding than of muscimol binding (Tang et al., 1993), though this observation has not been further explored. It is also known that only minor changes in the amino acid chains may alter the relative affinities of receptors for glycine and taurine, and there exist marked species differences and alterations during development (see > Sect. 5.2). The matter of the independent identity of taurine receptors is thus not yet settled.

4

Taurine Release

Basal release of taurine from neural cells can be mediated by different mechanisms (Saransaari and Oja, 1992): simple leakage through membranes (Saransaari and Oja, 1999d, 2006), flux through anionic Cl
channels (Saransaari and Oja, 1999b), and the reverse action of cell membrane carriers (Saransaari and Oja, 1999e, 2006). Under resting conditions the extracellular taurine concentration in the brain in vivo (Lerma et al., 1986; Mene´ndez et al., 1993; Segovia et al., 1997; Seki et al., 1999; Zielin´ska et al., 2002; Molchanova et al., 2004b; Ahmad et al., 2005) appears to be about one magnitude greater than that of the established neurotransmitter glutamate (Miele et al., 1996; Lai et al., 2000), but the interstitial concentrations of definitely nonneuroactive amino acids are even much higher (Lerma et al., 1986). Taurine release appears to be regulated and potentiated by many factors. The agonists of all ionotropic glutamate receptor classes have been shown to evoke taurine release from many different CNS preparations in vitro (Magnusson et al., 1991; Saransaari and Oja, 1991; McCaslin and Yu, 1992; Mene´ndez et al., 1993; Saransaari and Oja, 1994, 1997a, b) and in vivo (Jacobson and Hamberger, 1985; Sundstro¨m et al., 1995; Scheller et al., 2000a, b). Endogenous glutamate evokes an increase in the extracellular levels of taurine, acting through both NMDA and ()‐2‐amino‐3‐hydroxy‐5‐methylisoxazole‐4‐propionate (AMPA)/kainate receptors in the rat striatum (Segovia et al., 1997), AMPA receptors also being involved in the primary motor cortex (La Bella and Piccoli, 2000). The NMDA receptor antagonists block the NMDA‐evoked taurine release from mouse cerebral cortical slices in vitro (Saransaari and Oja, 1991, 2003b), in the rat hippocampus in vivo (Katoh et al., 1997), and in the rat spinal cord in vivo (Sundstro¨m et al., 1995). To the best of our knowledge we alone have studied the effects of metabotropic glutamate receptors on taurine release. The agonists of group I metabotropic glutamate receptors evoke taurine release, while the group II agonists are only slightly effective in the mouse hippocampus (Saransaari and Oja, 1999c). In slices from the brain stem, the agonists of groups I and II attenuate the Kþ‐evoked release in adult mice, and the agonists of group I and III in developing mice (Saransaari and Oja, 2006). NMDA receptor activation is thought to be mediated via the NO cascade (Scheller et al., 2000b), leading to the formation of cyclic GMP (cGMP) and the release of neurotransmitters and taurine (Guevara‐Guzman et al., 1994; Saransaari and Oja, 2002). Taurine release is indeed enhanced by nitric oxide donors from the mouse cerebral cortical neurons in culture (Chen et al., 1996), in hippocampal slices in vitro (Saransaari and Oja, 1999a), and in the rat striatum (Guevara‐Guzman et al., 1994) and the parietal cortex (Scheller et al., 2000b) in vivo. In keeping with this, the NO synthase blockers inhibit Kþ‐stimulated (Bo¨ckelmann et al., 1998) and NMDA‐mediated taurine release (Scheller et al., 2000b). The downstream cGMP also induces taurine release in the hippocampus (Saransaari and Oja, 2002). We here further refer to our relatively recent review on the roles of glutamate receptors and nitric oxide in taurine release (Oja and Saransaari, 2000). Adenosine stimulates cAMP‐mediated taurine release from LRM55 astroglial cells, apparently through adenosine A2 receptors (Madelian et al., 1988), and the release from the mouse (Saransaari and Oja, 2000, 2003a) and rabbit (Miyamoto and Miyamoto, 1999) hippocampus by the activation of adenosine

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A1 receptors. Furthermore, the adenosine transport inhibitors dipyradimole and nitrobenzylthioinosine significantly enhance taurine release from the rat hippocampus in vivo (Hada et al., 1996). In view of the possible neurotransmitter role of taurine, this depolarization‐evoked release is of significance. All experimental conditions that cause membrane depolarization enhance taurine release. On the other hand, membrane hyperpolarization shifts taurine transport from release toward uptake, resulting in seemingly diminished release (Lewin et al., 1994). Release of preloaded labeled taurine evoked by depolarizing concentrations of Kþ has been demonstrated in many neuronal preparations from the CNS (Kontro, 1979; Kontro and Oja, 1987b; Holopainen and Kontro, 1989; Oja et al., 1990a; Rogers et al., 1991; Oja and Saransaari, 1995; Zheng et al., 2000; Saransaari and Oja, 2006). However, when cell swelling is prevented, Kþ stimulation fails to evoke any release from astrocytes (Pasantes‐Morales and Schousboe, 1989; Pasantes‐Morales et al., 1990; Oja and Saransaari, 1992). The chemical identity of the released taurine has been confirmed by showing that potassium stimulation evokes release of endogenous taurine in brain tissue both in vitro and in vivo (Oja and Kontro, 1989; Sved and Curtis, 1993; Oja and Saransaari, 1995; Bo¨ckelmann et al., 1998; Colivicchi et al., 1998; Saransaari and Oja, 1998b; Estevez et al., 2000; Molchanova et al., 2004a). Also electrical stimulation evokes release of endogenous taurine from hippocampal, cerebral cortical, and cerebellar slices from the rat (Kubo et al., 1992), and transcranial stimulation, which activates descending motor fibers, elicits release in the rabbit lumbar spinal cord (Simpson et al., 1991). Furthermore, veratridine, which opens the voltage‐dependent Naþ channels, also stimulates taurine release from superfused slices from the rat substantia nigra (Della Corte et al., 1990) and the selective Naþ/Kþ‐ATPase inhibitor ouabain has the same effect in slices from the adult (Saransaari and Oja, 1998a) and developing mouse hippocampus (Saransaari and Oja, 1999e), human neuroblastoma cells (Basavappa et al., 1998), and rat cerebral cortex (Estevez et al., 2000). The stimulated release of taurine may result either from Ca2þ‐dependent emptying of synaptic vesicles or directly from cytoplasm (Molchanova et al., 2005). Calcium dependence is generally assumed to witness the exocytotic release of a neurotransmitter originating from the emptying of synaptic vesicles into the synaptic cleft. Various studies have yielded contradictory data on this matter. The Kþ‐stimulated release of taurine has been reported to be independent of external Ca2þ in cultured cerebellar astrocytes (Holopainen et al., 1985) and neurons (Rogers et al., 1991), goldfish cerebellar slices (Lucchi et al., 1994; Rosati et al., 1995), and apparently also in the rat hippocampus (Solı´s et al., 1986) and spinal cord in vivo (Sundstro¨m et al., 1995). In contrast, the Kþ‐stimulated release has been found to be Ca2þ‐dependent in synaptosomes from the rat cerebral cortex (Kamisaki et al., 1996b) and the rat substantia nigra in vivo (Garcı´a Dopico et al., 2004). Moreover, the Ca2þ‐dependent component could be extracted by computer analysis from the total Kþ‐evoked release of taurine from cerebral cortical synaptosomes (Kontro, 1979), and a part of the evoked taurine release from cortical synaptosomes was also recently estimated to be exocytotic in nature (Tuz et al., 2004). In accord, the Kþ‐stimulated release in hippocampal and brain stem slices is partially Ca2þ dependent in both developing and adult mice (Saransaari and Oja, 1998a, b, 1999e, 2006). Ca2þ dependence may depend on the methods used to elicit the release and intensity of stimulus, and the release of Ca2þ from intracellular stores may support taurine release (Mene´ndez et al., 1993). The question of the Ca2þ dependency of taurine release is thus not yet settled.

5

Neuromodulatory Functions

5.1 Interactions with GABAergic Systems As a close structural analog taurine could interfere with the uptake and release of GABA (see > Sects. 2.3 and > 4). In synaptic structures, it displaces GABA (Malminen and Kontro, 1986, 1987) and muscimol (Egbuta and Griffiths, 1987) binding to the GABA–benzodiazepine receptor complex and interferes with ligand binding to its benzodiazepine site (Iwata et al., 1984; Medina and De Robertis, 1984; Malminen and Kontro, 1986, 1987) allosterically (Quinn and Miller, 1992). Taurine activates GABAA receptors (Horikoshi et al., 1988; Zhu and Vicini, 1997; del Olmo et al., 2000; Barakat et al., 2002), but less effectively than GABA itself. It binds to the GABAA receptors in hippocampal and cortical neurons and prevents b‐amyloid

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peptide neurotoxicity (Paula‐Lima et al., 2005). Taurine also attenuates 1‐methyl‐4‐phenylpyridinium (MPPþ) neurotoxicity in coronal slices from the rat brain by means of activation of GABAA receptors (O’Byrne and Tipton, 2000). The GABAA antagonists bicuculline and picrotoxin block taurine effects on GABAA receptors solubilized from the rat brain (Malminen and Kontro, 1987) and on these receptors in the mitral and tufted cells in the rat main olfactory bulb (Belluzzi et al., 2004). At the GABAA receptor complex taurine is a full agonist of GABA (Quinn and Harris, 1995), but preferentially only in the b2 subunit‐rich areas of GABAA receptors, enriched in the dentate gyrus, substantia nigra, cerebellar molecular layer, median thalamic nuclei, and hippocampal CA3 field (Bureau and Olsen, 1991). Differences in taurine sensitivity may stem from the different subunit composition of GABAA receptors, although Rabe and his coworkers (2000) found no differences in the regional efficacies of taurine and GABA. In the nucleus accumbens of young rats (Jiang et al., 2004) and in cultured cerebellar granule cells (Wahl et al., 1994), taurine is only a partial agonist. Treatment of cultured cerebellar granule cells with taurine induces low‐ affinity GABA receptors (Abraham and Schousboe, 1989), but chronic supplementation to mice induces a downregulation of expression of the GABAA receptor b subunit (the key subunit present in virtually all GABAA receptors) (El Idrissi, 2006). On the other hand, glutamate decarboxylase‐positive neurons increase in number in the cerebral cortex of taurine‐fed mice (Levinskaya et al., 2006). In some instances taurine has been reported to affect GABAB receptors (Kontro and Oja, 1990; Kontro et al., 1990). It has proved an even more potent displacer at GABAB than at GABAA receptors in rat brain membranes (Malminen and Kontro, 1986). In the frog olfactory bulb in vivo, taurine suppresses the spontaneous firing of mitral cells and acts mainly on GABAB receptors (Chaput et al., 2004) by increasing the signal‐ to‐noise ratio. In the rat, taurine has likewise suppressed olfactory nerve‐evoked monosynaptic responses of mitral and tufted cells and blocked chloride conductance, probably owing to the GABAB receptor‐mediated inhibition of glutamate release (Belluzzi et al., 2004). Taurine may also regulate GABA release from rat brain olfactory bulb synaptosomes via activation of presynaptic GABAB receptors (Kamisaki et al., 1993, 1996a). The paucity of reports may signify that taurine affects GABAB receptors only in limited cases.

5.2 Interactions with Glycinergic Systems Taurine interacts with glycinergic systems only at the receptor level, since it is not a preferred substrate for glycine transporters. It displaces glycine from its strychnine‐sensitive binding sites in the mouse brain stem (Kontro and Oja, 1987c) and acts as an agonist at glycine receptors in Xenopus oocytes injected with mouse brain mRNA (Horikoshi et al., 1988), in the rat supraoptic magnocellular neurons (Hussy et al., 1997), and in dopaminergic neurons in the ventral tegmental area in young rats (Ye et al., 2004). In mammalian receptors, glycine is more effective than taurine, but in Hydra vulgaris glycine and taurine are almost equipotent (Pierobon et al., 2001). Taurine in some instances affects both GABAA and glycine receptors simultaneously. For example, it controls hormone release from the rat neurohypophysis (Song and Hatton, 2003) and prevents the ammonia‐ induced accumulation of cGMP in the rat striatum in this manner (Hilgier et al., 2005). At low concentrations taurine activates glycine receptors in CA1 neurons in the immature rat hippocampus, whereas at high concentrations both glycine and GABA receptors are activated (Wu and Xu, 2003). In mice, the target of taurine shifts from glycine receptors to GABA receptors during postnatal development (Yoshida et al., 2004). The similar effects of taurine and glycine on Cl
conductance have already been dealt with in > Sect. 3.1. Taurine may be an important endogenous ligand for glycine receptors in certain parts of the CNS. For instance, the glycine‐receptor clusters and the glial fibrillary acidic protein‐positive astroglial processes, which contain high levels of taurine in the rat supraoptic nucleus, are anatomically associated (Deleuze et al., 2005). The activation by taurine of glycine receptors in neurohypophysial nerve terminals is thought to participate in the osmoregulation of vasopressin secretion (Hussy et al., 2001). It acts mainly on nonsynaptic glycine receptors in the ventral tegmental area, and their dopaminergic neurons could be exposed tonically to taurine (Wang et al., 2005).

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Taurine has been shown to inhibit the glycine responses of oocytes injected with rat a1 subunits of the glycine receptor or spinal cord poly(A)þ RNA (Schmieden et al., 1989). It is a full agonist of the glycine receptors in the ventral tegmental area (Wang et al., 2005). Taurine more efficiently gates glycine a1 receptors than a2 receptors, but glycine is more effective than taurine in both receptor subtypes (Schmieden et al., 1992). Human and rat a2 and a3 glycine receptors display rather low responses to taurine, and the amino acid 167 in the a2 subunit is essential for taurine effects (Kuhse et al., 1990a, b). On the other hand in the mammalian a1 subunits, isoleucine in position 111 and alanine in 212 have been considered important determinants of taurine activation, and valine in these positions in a2 may reduce the efficacy of taurine (Schmieden et al., 1992). The glycine receptor a1 subunit of the zebrafish (aZ1) is very similar to the rat a1 receptor in both length and amino acid residue composition, but the homomeric aZ1 receptor exhibits exceptionally high sensitivity for taurine, almost comparable to glycine sensitivity (David‐Watine et al., 1999). It contains valine instead of isoleucine in position 111 and the authors infer that the natural substitution Ile111Val may not be a critical determinant for taurine selectivity. Sequence differences in the immediate vicinity of Val111 may rather underlie the high efficacy of taurine. In other experiments, taurine has also acted as a full agonist in aZ2 glycine receptors in spite of the presence of Val111 and Val212 (Imboden et al., 2001). The more recently cloned homomeric a4 subunit from chicks containing valines in the equivalent positions is also potently activated by taurine (Harvey et al., 2000). Two separate point mutations at the same base pair in the gene encoding the human glycine receptor a1 subunit, resulting in arginine at position 271 being substituted by leucine (R271L) or glutamine (R271Q), have been observed to convert taurine from a full agonist to a competitive antagonist (Laube et al., 1995; Rajendra et al., 1995). The same changes occur with the Y279C and K276E mutations (tyrosine to cysteine and lysine to glutamate, respectively) (Lynch et al., 1997). It is assumed that these mutations either simply disconnect the binding site of taurine from the Cl
channel activation site or selectively disrupt a common agonist recognition subsite and thereby unmask the antagonist subsite for taurine (Laube et al., 1995; Rajendra et al., 1995; Schmieden and Betz, 1995). Furthermore, three mutations, R218Q, V260M, and Q266H (arginine to glutamine, valine to methionine, and glutamine to histidine, respectively), in the a1 subunit of the human glycine receptor GlyRA1, which cause hyperekplexia, greatly reduce Cl
currents elicited by taurine (Castaldo et al., 2004).

5.3 Interactions with Other Transmitter Systems There are only scattered data on the effects of taurine on other neurotransmitter systems. It modulates noradrenaline uptake and release in rat cerebral cortical slices (Kontro et al., 1984), interferes with the binding of labeled spiperone (serotonin 5‐HT2A and dopamine D2 receptor antagonist) to cerebral cortical membranes (Kontro and Oja, 1986), and interacts with dopaminergic neurotransmission in the striatum (Kontro, 1987; Kontro and Oja, 1988a). The age‐related decline in striatal taurine is correlated with a loss of dopaminergic markers (Dawson et al., 1999). Intranigrally injected taurine modulates striatal dopaminergic transmission (O’Neill, 1986) and reduces extracellular dopamine (Leviel et al., 1979; Ruotsalainen et al., 1996). Intracerebroventricular (Ahtee and Vahala, 1985) and intraperitoneal (Salima¨ki et al., 2003) taurine and taurine administered directly into the striatum (Ruotsalainen et al., 1998) have significantly increased extracellular dopamine (Ruotsalainen and Ahtee, 1996). At variance, in other studies direct administration into the rat striatum did not markedly affect extracellular dopamine, but reduced extracellular dopamine metabolite dihydroxyphenylacetic acid and accentuated its NMDA‐induced decrease (Anderzhanova et al., 2001, 2006). Taurine inhibits the synthesis and release of serotonin in rat rostral, but not in caudal, rhombencephalic raphe cells (Becquet et al., 1993). It also affects the binding of phencyclidine (which blocks NMDA receptor activation) in the mouse cerebral cortex (Saransaari and Oja, 1993). Finally, taurine reduces Kþ‐stimulated adenosine release in slices from the hippocampus in the developing but not in adult mice (Saransaari and Oja, 2003a).

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5.4 Effects on Calcium Levels A neuromodulator could affect calcium homeostasis, and thus, for example, modify neurotransmitter release and synthesis. Indeed, taurine has inhibited NMDA‐evoked Ca2þ accumulation in brain slices (Lehmann et al., 1984), attenuated Ca2þ influx into slices from developing mice (Kontro and Oja, 1988b), and protected neurons from glutamate‐induced excitotoxicity (Tang et al., 1996), this by preventing or reducing the glutamate‐induced elevation of intracellular Ca2þ (Chen et al., 2001). Taurine also inhibits the reverse mode of the Naþ/Ca2þ exchangers (Wu et al., 2000) and protects cultured rat astrocytes against reperfusion injury (Matsuda et al., 1996). It may not directly affect the rate of Ca2þ uptake but rather the duration of the maximal response to glutamate (El Idrissi and Trenkner, 1999). Taurine also inhibits the glutamate‐induced release of Ca2þ from the internal pools (Wu et al., 2000) and regulates cytoplasmic and mitochondrial calcium homeostasis (El Idrissi and Trenkner, 2003). In addition to this, taurine inhibits the glutamate‐induced Ca2þ influx through L‐, P/Q‐, and N‐types of voltage‐gated calcium channels and the NMDA receptor‐governed calcium channel in whole‐brain primary neuronal cultures (Wu et al., 2005). On the other hand, it has been shown to increase the accumulation of 45Ca2þ in mitochondria of the rat cerebral cortex, being thus able to attenuate the cytosolic free Ca2þ concentration, which, in turn, inhibits specific protein phosphorylation and phosphoinositide turnover (Li and Lombardini, 1991). Taurine efficiently counteracts the glutamate‐induced Ca2þ uptake in cerebellar granule cells (El Idrissi et al., 1998; Trenkner et al., 1998). In brain slices, taurine forestalls cell damage evoked by overactivation of ionotropic glutamate receptors (Zielin´ska et al., 2003). Overactivation of NMDA receptors leads to mitochondrial damage associated with Ca2þ influx, which results in the generation of free radicals, including superoxide. It is thought that a part of the neuroprotective action of taurine depends on its antioxidant properties (Schaffer et al., 2003). Taurine has been shown to antagonize the Ca2þ overload induced by glutamate and hypoxia in cultured rat hippocampal neurons (Zhao et al., 1999). On the whole, taurine may have an essential role in the modulation of intracellular calcium homeostasis in both normal and cell‐damaging conditions (Chen et al., 2001; Foos and Wu, 2002).

6

Conclusions and Perspectives

Present knowledge of the possible roles of taurine as a neurotransmitter or neuromodulator remains incomplete. There are ample amounts of taurine in nerve cells and synaptic structures for both functions, which can be terminated by an efficient reuptake into neurons or glia. Taurine is released by depolarization, though the release mechanisms have not been thoroughly characterized, the Ca2þ dependency of the stimulated release being the most essential question to settle. It increases membrane chloride conductance, which effect is receptor‐mediated. However, more investigation is still needed to prove or disprove the existence of taurine receptors. Binding studies and isolation and possible cloning of the binding sites are essential, in addition to investigations with other animal species than mammals. These studies should be supplemented with analyses of the electrical responses of the receptors and tissue preparations to taurine. Taurine is not a neuromodulator in the classical sense, as it has actions of its own and does not of itself influence the functions of established neurotransmitters. It is still open whether or not all taurine actions in neurotransmission are attributable to its interactions with GABA and glycine receptors. In particular, the cloning of different glycine receptors from various sources and their mutations, which alter the responses to taurine, glycine, and GABA, would suggest the existence of independent taurine receptors. It may be noted that the efficacy of taurine at glycine receptors appears to be greater in developing animals and in animal species other than mammals. Even in mammals some effects in certain brain areas could be mediated by receptors specific for taurine.

Acknowledgments Financial support from the competitive research funding of the Pirkanmaa Hospital District (grants 9F051, 9F068, 9G051, and 9G068) is gratefully acknowledged.

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References Abraham JH, Schousboe A. 1989. Effects of taurine on cell morphology and expression of low‐affinity GABA receptors in cultured cerebellar granule cells. Neurochem Res 14: 1031-1038. Ahmad S, Fowler LJ, Whitton PS. 2005. Lamotrigine, carbamazepine and phenytoin differentially alter extracellular levels of 5‐hydroxytryptamine, dopamine and amino acids. Epilepsy Res 63: 141-149. Ahtee L, Vahala ML. 1985. Taurine and its derivatives alter brain dopamine metabolism similarly to GABA in mice and rats. Prog Clin Biol Res 179: 331-341. Ament ME, Geggel HS, Heckenlively JR, Martin DA, Kopple JD. 1986. Taurine supplementation in infants receiving long‐term total parenteral nutrition. J Am Coll Nutr 5: 127-135. Anderzhanova E, Rayevsky KS, Saransaari P, Riitamaa E, Oja SS. 2001. Effects of sydnocarb and D‐amphetamine on the extracellular levels of amino acids in the rat caudate‐ putamen. Eur J Pharmacol 428: 87-95. Anderzhanova E, Saransaari P, Oja SS. 2006. Neuroprotective mechanisms of taurine in vivo. Adv Exp Med Biol 583: 377-387. Barakat L, Wang D, Bordey A. 2002. Carrier‐mediated uptake and release from Bergman glia in rat cerebellar slices. J Physiol 541: 753-767. Basavappa S, Mobasheri A, Errington R, Huang CC, Al‐Adawi S, et al. 1998. Inhibition of Naþ, Kþ‐ATPase activates swelling‐induced taurine efflux in a human neuroblastoma cell line. J Cell Physiol 174: 145-153. Beckman ML, Quick MW. 1998. Neurotransmitter transporters: Regulators of function and functional regulation. J Membr Biol 164: 1-10. Becquet D, Hery M, Francois‐Bellan AM, Giraud P, Deprez P, et al. 1993. Glutamate, GABA, glycine and taurine modulate serotonin synthesis and release in rostral and caudal rhombencephalic raphe cells in primary cell cultures. Neurochem Int 23: 269-283. Beetsch JW, Olson JE. 1996. Hyperosmotic exposure alters total taurine quantity and cellular transport in rat astrocyte cultures. Biochim Biophys Acta 1290: 141-148. Beetsch JW, Olson JE. 1998. Taurine synthesis and cysteine metabolism in cultured rat astrocytes: Effects of hyperosmotic exposure. Am J Physiol 274: C866-C874. Belluzzi D, Puopolo M, Benedusi M, Kratskin I. 2004. Selective neuroinhibitory effects of taurine in slices of rat main olfactory bulb. Neuroscience 124: 929-944. Bitoun M, Tappaz M. 2000. Taurine down‐regulates basal and osmolarity‐induced gene expression of its transporter, but not the gene expression of its biosynthetic enzymes, in astrocyte primary cultures. J Neurochem 75: 919-924.

Bo¨ckelmann R, Reiser M, Wolf G. 1998. Potassium‐stimulated taurine release and nitric oxide synthase activity during quinolinic acid lesion of the rat striatum. Neurochem Res 23: 469-475. Borden LA, Smith KE, Vaysse PJ, Gustafson EL, Weinshank RL, et al. 1995. Re‐evaluation of GABA transport in neuronal and glial cell cultures: Correlation of pharmacology and mRNA localization. Receptors Channels 3: 129-146. Braghiroli D, Di Bella M, Zanoli P, Truzzi C, Baraldi M. 1990. Rigid analogs of taurine as potential taurine antagonists. Farmaco 45: 631-645. Bureau MH, Olsen RW. 1991. Taurine acts on a subclass of GABAA receptors in mammalian brain in vitro. Eur J Pharmacol 207: 9-16. Castaldo P, Stefanoni P, Miceli F, Coppola G, Miraglia del Giudice E, et al. 2004. A novel hyperekplexia‐causing mutation in the pre‐transmembrane segment 1 of the human glycine receptor a1 subunit reduces membrane expression and impairs gating by agonists. J Biol Chem 279: 25598-25604. Chaput MA, Palouzier‐Paulignan B, Delaleu JC, Duchamp‐ Viret P. 2004. Taurine action on mitral cell activity in the frog olfactory bulb in vivo. Chem Senses 29: 83-91. Chen DZ, Ohkuma S, Kuriyama K. 1996. Characteristics of nitric oxide‐evoked [3H]taurine release from cerebral cortical neurons. Neurochem Int 28: 601-607. Chen WQ, Jin H, Nguyen M, Carr J, Lee YJ, et al. 2001. Role of taurine in regulation of intracellular calcium level and neuroprotective function in cultured neurons. J Neurosci Res 66: 612-619. Chesney RW, Gusowski N, Dabbaugh S. 1985. Renal cortex taurine regulates the adaptive response to altered dietary intake of sulfur amino acids. J Clin Invest 76: 2213-2221. Colivicchi MA, Bianchi L, Boloam JP, Galeffi F, Frosini M, et al. 1998. The in vivo release of taurine in the striatonigral pathway. Adv Exp Med Biol 442: 363-370. Coloso RM, Hirschberger LL, Dominy JE, Lee JI, Stipanuk MH. 2006. Cysteamine dioxygenase: Evidence for the physiological conversion of cysteamine to hypotaurine in rat and mouse tissues. Adv Exp Med Biol 583: 25-35. Cutler RW, Dudzinski DS. 1974. Regional changes in amino acid content in developing rat brain. J Neurochem 23: 1005-1009. David‐ Watine B, Goblet C, de Saint Jan D, Fucile S, Devignot V, et al. 1999. Cloning, expression and electrophysiological characterization of glycine receptor a subunit from zebrafish. Neuroscience 90: 303-317. Dawson R Jr, Pelleymounter MA, Cullen MJ, Gollub M, Liu S. 1999. An age‐related decline in striatal taurine is correlated with a loss of dopaminergic markers. Brain Res Bull 48: 319-324.

335

336

13

Taurine in neurotransmission

Decavel C, Hatton GI. 1995. Taurine immunoreactivity in the rat supraoptic nucleus: Prominent localization in glial cells. J Comp Neurol 354: 13-26. Deleuze C, Alonso G, Lefevre IA, Duvoid‐Guillou A, Hussy N. 2005. Extrasynaptic localization of glycine receptors in the rat supraoptic nucleus: Further evidence for their involvement in glia‐to‐neuron communication. Neuroscience 133: 175-183. Della Corte L, Bolam JP, Clarke DJ, Parry DM, Smith AD. 1990. Sites of [3H]taurine uptake in the rat substantia nigra in relation to the release of taurine from the nigrostriatal pathway. Eur J Neurosci 2: 50-61. del Olmo N, Bustamante J, Martı´n del Rı´o R, Solı´s JM. 2000. Taurine activates GABAA but not GABAB receptors in rat hippocampal CA1 area. Brain Res 864: 298-307. Didier A, Ottersen OP, Storm‐Mathisen J. 1994. Differential subcellular distribution of glutamate and taurine in primary olfactory neurons. Neuroreport 6: 145-148. Egbuta JO, Griffiths R. 1987. Binding of [3H]muscimol to calf cerebrocortical synaptic membranes and the effects of sulphur‐containing convulsant and non‐convulsant compounds. Neurochem Res 12: 589-595. El Idrissi A. 2006. Taurine and brain excitability. Adv Exp Biol Med 583: 315-322. El Idrissi A, Trenkner E. 1999. Growth factors and taurine protect against excitotoxicity by stabilizing calcium homeostasis and energy metabolism. J Neurosci 19: 9459-9468. El Idrissi A, Trenkner E. 2003. Taurine regulates mitochondrial calcium homeostasis. Adv Exp Med Biol 526: 527-536. El Idrissi A, Harris C, Trenkner E. 1998. Taurine modulates glutamate and growth factors‐mediated signaling mechanisms. Adv Exp Med Biol 442: 385-396. Estevez AY, Song D, Phillis JW, O’Regan MH. 2000. Effects of the anion channel blocker DIDS on ouabain‐ and high Kþ‐ induced release of amino acids from the rat cerebral cortex. Brain Res Bull 52: 45-50. Fellman JH, Roth ES. 1985. The biological oxidation of hypotaurine to taurine: Hypotaurine as an antioxidant. Prog Clin Biol Res 179: 71-82. Foos TM, Wu JY. 2002. The role of taurine in the central nervous system and the modulation of intracellular calcium homeostasis. Neurochem Res 27: 21-26. Frosini M, Sesti C, Dragoni S, Valoti M, Palmi M, et al. 2003. Interactions of taurine and structurally related analogues with the GABAergic system and taurine‐binding sites of rabbit brain. Br J Pharmacol 138: 1163-1171. Fujita T, Shimada A, Wada M, Miyakawa S, Yamamoto A. 2006. Functional expression of taurine transporter and its up-regulation in developing neurons from mouse cerebral cortex. Pharm Res 23: 689-696.

Garcı´a Dopico J, Perdomo Dı´az J, Alonso TJ, Gonza´lez Herna´ndez T, Castro Fuentes R, et al. 2004. Extracellular taurine in the substantia nigra: Taurine–glutamate interaction. J Neurosci Res 76: 528-538. Geggel HS, Ament ME, Heckenlively JR, Martin DA, Kopple JD. 1985. Nutritional requirement for taurine in patients receiving long‐term parenteral nutrition. N Engl J Med 312: 142-146. Godfrey DA, Farms WB, Godfrey TG, Mikesell NL, Liu J. 2000. Amino acid concentrations in rat cochlear nucleus and superior olive. Hearing Res 150: 189-205. Gragera RR, Muniz E, De Esteban G, Alonso MJ, Martı´nez‐ Rodrı´guez R. 1995. Immunochemical demonstration of taurine in the rat cerebellar cortex. Evidence for its location within mossy fibers and Golgi axons. J Hirnforsch 36: 269-276. Guevara‐Guzman R, Emson PC, Kendrick KM. 1994. Modulation of in vivo striatal transmitter release by nitric oxide and cyclic GMP. J Neurochem 62: 807-810. Guidotti A, Badiani G, Pepeu G. 1972. Taurine distribution in cat brain. J Neurochem 19: 431-435. Hada J, Kaku T, Morimoto K, Hayashi Y, Nagai K. 1996. Adenosine transport inhibitors enhance high Kþ‐evoked taurine release from rat hippocampus. Eur J Pharmacol 305: 101-107. Hanretta AT, Lombardini JB. 1987. Is taurine a hypothalamic neurotransmitter?: A model of the differential uptake and compartmentalization of taurine by neuronal and glial cell particles from the rat hypothalamus. Brain Res 434: 167-201. Harvey RJ, Schmieded V, von Holst A, Laube B, Rohrer H, et al. 2000. Glycine receptors containing the a4 subunit in the embryonic sympathetic nervous system, spinal cord and male genital ridge. Eur J Neurosci 12: 994-1001. Ha¨user MA, Yung WH, Lacey MG. 1992. Taurine and glycine activate the same Cl
conductance in substantia nigra dopamine neurons. Brain Res 571: 103-108. Hilgier W, Oja SS, Saransaari P, Albrecht J. 2005. Taurine prevents ammonia‐induced accumulation of cyclic AMP in rat striatum by interaction with GABAA and glycine receptors. Brain Res 1043: 242-246. Holopainen I, Kontro P. 1989. Uptake and release of glycine in cerebellar granule cells and astrocytes in primary culture: The potassium‐stimulated release from granule cells is Ca‐ dependent. J Neurosci Res 24: 374-383. Holopainen I, Kontro P, Frey HJ, Oja SS. 1983. Taurine, hypotaurine, and GABA uptake by cultured neuroblastoma cells. J Neurosci Res 10: 83-92. Holopainen I, Kontro P, Oja SS. 1985. Release of preloaded taurine and hypotaurine from astrocytes in primary culture: Stimulation by calcium‐free media. Neurochem Res 10: 123-131.

Taurine in neurotransmission Horikoshi T, Asanuma A, Yanagisawa K, Anzai K, Goto S. 1988. Taurine and b‐alanine act on both GABA and glycine receptors in Xenopus oocyte injected with mouse brain messenger RNA. Brain Res 464: 97-105. Hussy N, Bre`s V, Rochette M, Duvoid A, Alonso G, et al. 2001. Osmoregulation of vasopressin secretion via activation of neurohypophysial nerve terminals glycine receptors by glial taurine. J Neurosci 21: 7110-7116. Hussy N, Deleuze C, Pantaloni A, Desarme´nien MG, Moos F. 1997. Agonist action of taurine on glycine receptors in rat supraoptic magnocellular neurons: Possible role in osmoregulation. J Physiol 502: 609-621. Huxtable RJ. 1986. Taurine and the oxidative metabolism of cysteine. Biochemistry of Sulfur. New York: Plenum Press; pp. 121–198. Huxtable RJ. 1989. Taurine in the central nervous system and the mammalian actions of taurine. Prog Neurobiol 32: 471-533. Imaki H, Moretz RC, Wisniewski HN, Neuringer M, Sturman JA. 1987. Retinal degeneration in three‐month‐old rhesus monkey infants fed a taurine‐free human infant formula. J Neurosci Res 18: 602-614. Imaki H, Moretz RC, Wisniewski HM, Sturman JA. 1986. Feline maternal taurine deficiency: Effects on retina and tapetum of the offspring. Dev Neurosci 8: 160-181. Imboden M, De Saint Jan D, Leulier F, Korn H, Coblet C, et al. 2001. Isolation and characterization of an a2‐type zebrafish glycine receptor subunit. Neuroscience 103: 799-810. Iwata H, Nakayama K, Matsuda T, Baba A. 1984. Effect of taurine on a benzodiazepine‐GABA‐chloride ionophore receptor complex in rat brain membranes. Neurochem Res 9: 535-544. Jacobson I, Hamberger A. 1985. Kainic acid‐induced changes of extracellular amino acid levels, evoked potentials and EEG activity in the rabbit olfactory bulb. Brain Res 348: 289-296. Jiang Z, Krnjevic´ K, Wang F, Ye JH. 2004.Taurine activates strychnine‐sensitive glycine receptors in neurons freshly isolated from nucleus accumbens of young rats. J Neurophysiol 91: 248-257. Kamisaki Y, Maeda K, Ishimura M, Omura H, Itoh T. 1993. Effects of taurine on depolarization‐evoked release of amino acids from rat cortical synaptosomes. Brain Res 627: 181-185. Kamisaki Y, Wada K, Nakamoto K, Itoh T. 1996a. Effect of taurine on GABA release from synaptosomes of rat olfactory bulb. Amino Acids 10: 49-57. Kamisaki Y, Wada K, akamoto NK, Itoh T. 1996b. Release of taurine and its effects on release of neurotransmitter amino acids in rat cerebral cortex. Adv Exp Med Biol 403: 445-454. Kang YS. 2000. Taurine transport mechanism through the blood–brain barrier in spontaneously hypertensive rats. Adv Exp Med Biol 483: 321-324.

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Katoh H, Sima K, Nawashiro H, Wada K, Chigasaki H. 1997. The effect of MK‐801 on extracellular neuroactive amino acids in hippocampus after closed head injury followed by hypoxia in rats. Brain Res 758: 153-162. Kontro P. 1979. Components of taurine efflux in rat brain synaptosomes. Neuroscience 4: 1745-1749. Kontro P. 1987. Interactions of taurine and dopamine in the striatum. Adv Exp Med Biol 217: 347-355. Kontro P, Oja SS. 1978. Taurine uptake by rat brain synaptosomes. J Neurochem 30: 1297-1304. Kontro P, Oja SS. 1983. Sodium‐independent taurine binding to brain synaptic membranes. Cell Mol Neurobiol 3: 183-187. Kontro P, Oja SS. 1985a. Hypotaurine oxidation by mouse liver tissue. Prog Clin Biol Res 179: 83-90. Kontro P, Oja SS. 1985b. Properties of sodium‐independent taurine binding to brain synaptic membranes. Prog Clin Biol Res 179: 249-259. Kontro P, Oja SS. 1986. Taurine interferes with spiperone binding in the striatum. Neuroscience 19: 1007-1010. Kontro P, Oja SS. 1987a. Taurine and GABA binding in mouse brain: Effects of freezing, washing and Triton X‐100 treatment on membranes. Int J Neurosci 32: 881-889. Kontro P, Oja SS. 1987b. Taurine and GABA release from mouse cerebral cortex slices: Potassium stimulation releases more taurine than GABA from developing brain. Dev Brain Res 37: 277-291. Kontro P, Oja SS. 1987c. Glycinergic systems in the brain stem of developing and adult mice: Effects of taurine. Int J Dev Neurosci 5: 461-470. Kontro P, Oja SS. 1987d. Co‐operativity in sodium‐independent taurine binding to brain membranes in the mouse. Neuroscience 23: 567-570. Kontro P, Oja SS. 1988a. Release of taurine, GABA and dopamine from rat striatal slices: Mutual interactions and developmental aspects. Neuroscience 24: v-58. Kontro P, Oja SS. 1988b. Effects of taurine on the influx and efflux of calcium in brain slices of adult and developing mice. Int J Neurosci 38: 103-109. Kontro P, Oja SS. 1990. Interactions of taurine with GABAB binding sites in mouse brain. Neuropharmacology 29: 243-247. Kontro P, Korpi ER, Oja OS, Oja SS. 1984. Modulation of noradrenaline uptake and release by taurine in rat cerebral slices. Neuroscience 13: 663-666. Kontro P, Korpi ER, Oja SS. 1990. Taurine interacts with GABAA and GABAB receptors in the brain. Prog Clin Biol Res 351: 83-94. Kontro P, Marnela KM, Oja SS. 1980. Free amino acids in the synaptosome and synaptic vesicle fractions of different bovine brain areas. Brain Res 184: 129-141.

337

338

13

Taurine in neurotransmission

Krishtal OA, Osipchuk Yu V, Vrublevsky SV. 1988. Properties of glycine‐activated conductances in rat brain neurons. Neurosci Lett 84: 271-276. Kubo T, Takano A, Tokushige N, Miyata N, Sato M, et al. 1992. Electrical stimulation‐evoked release of endogenous taurine from slices of the hippocampus, cerebral cortex and cerebellum of the rat. J Pharmacobiodyn 15: 519-525. Kudo Y, Akiyoshi E, Akagi H. 1988. Identification of two taurine receptor subtypes on the primary afferent terminal of frog spinal cord. Br J Pharmacol 94: 1051-1056. Kuhse J, Schmieden V, Betz H. 1990a. Identification and functional expression of a novel ligand binding subunit of the inhibitory glycine receptor. J Biol Chem 265: 2231722320. Kuhse J, Schmieden V, Betz H. 1990b. A single amino acid exchange alters the pharmacology of neonatal rat glycine receptor subunit. Neuron 5: 867-873. Kulanthaivel P, Cool DR, Ramamoorthy S, Mahesh VB, Leibach FH, et al. 1991. Transport of taurine and its regulation by protein kinase C in the JAR human placental choriocarcinoma cell line. Biochem J 277: 53-58. La Bella V, Piccoli F. 2000. Differential effect of b‐N‐oxalylamino‐L‐alanine, the Lathyrus sativus neurotoxin, and ()‐a‐amino‐3‐hydroxy‐5‐methylisoxazole‐4‐propionate on the excitatory amino acid and taurine levels in the brain of freely moving rat. Neurochem Int 36: 523-530. La¨hdesma¨ki P, Oja SS. 1973. On the mechanism of taurine transport at brain cell membranes. J Neurochem 20: 1411-1417. Lai YJ, Shen EY, Pan WHT. 2000. Effects of ascorbate in microdialysis perfusion medium on the extracellular basal concentration of glutamate in rat’s striatum. Neurosci Lett 279: 145-148. Lake N, Orlowski J. 1996. Cellular studies of the taurine transporter. Adv Exp Med Biol 403: 371-376. Lallemand F, De Witte P. 2004. Taurine concentration in the brain and in the plasma following intraperitoneal injections. Amino Acids 26: 111-116. Lane JD, Smith JE, Hall PV, Campbell RL. 1978. Distribution of taurine and putative amino acid neurotransmitters in eight areas of the canine lumbar spinal cord. Brain Res 152: 386-390. Laube B, Langosch D, Betz H, Schmieden V. 1995. Hyperekplexia mutations of the glycine receptor unmask the inhibitory subsite for b‐amino acids. Neuroreport 6: 897-900. Lehmann A, Hagberg H, Hamberger A. 1984. A role for taurine in the maintenance of homeostasis in the central nervous system during hyperexcitation? Neurosci Lett 52: 341-346. Lerma J, Herranz AS, Herreras O, Abraira V, Martı´n del Rı´o R. 1986. In vivo determination of extracellular concentration of amino acids in the rat hippocampus. A method based on

brain dialysis and computerized analysis. Brain Res 384: 145-155. Leviel V, Che´ramy A, Nieoullon A, Glowinski J. 1979. Symmetric bilateral changes in dopamine release from the caudate nuclei of the cat induced by unilateral nigral application of glycine and GABA‐related compounds. Brain Res 175: 259-270. Levinskaya N, Trenkner E, El Idrissi A. 2006. Increased GAD‐ positive neurons in the cortex of taurine‐fed mice. Adv Exp Med Biol 583: 411-417. Lewin L, Rassin DK, Sellstro¨m A˚. 1994. Net taurine transport and its inhibition by a taurine antagonist. Neurochem Res 19: 347-352. Li YP, Lombardini JB. 1991. Inhibition by taurine of the phosphorylation of specific synaptosomal proteins in the rat cortex: Effects of taurine on the stimulation of calcium uptake in mitochondria and inhibition of phosphoinositide turnover. Brain Res 553: 89-96. Li H, Godfrey DA, Rubin AM. 1994. Quantitative distribution of amino acids in the rat vestibular nuclei. J Vest Res 4: 437-452. Li H, Godfrey TG, Godfrey DA, Rubin AM. 1996. Quantitative changes of amino acid distributions in the rat vestibular nuclear complex after unilateral vestibular ganglionectomy. J Neurochem 66: 1550-1564. Linne ML, Jalonen TO, Saransaari P, Oja SS. 1996. Taurine‐ induced single‐channel currents in cultured rat cerebellar granule cells. Adv Exp Med Biol 403: 455-462. Liu QR, Lo´pez‐Corcuera B, Mandiyan S, Nelson H, Nelson N. 1993. Molecular characterization of four pharmacologically distinct g‐aminobutyric acid transporters in mouse brain. J Biol Chem 268: 2106-2112. Liu QR, Lo´pez‐Corcuera B, Nelson H, Mandiyan S, Nelson N. 1992. Cloning and expression of cDNA encoding the transporter of taurine and b‐alanine in mouse brain. Proc Natl Acad Sci USA 89: 12145-12149. Lucchi R, Poli A, Traversa U, Barnabei O. 1994. Functional adenosine A1 receptors in goldfish brain: Regional distribution and inhibition of Kþ‐evoked glutamate release from cerebellar slices. Neuroscience 58: 237-243. Lynch JW, Rajendra S, Pierce KD, Handford CA, Barry PH, et al. 1997. Identification of intracellular and extracellular domains mediating signal transduction in the inhibitory glycine receptor chloride channel. EMBO J 16: 110-120. Maar T, Mora´n J, Schousboe A, Pasantes‐Morales H. 1995. Taurine deficiency in dissociated mouse cerebellar neurons affects neuronal migration. Int J Dev Neurosci 13: 491-502. Madelian V, Silliman S, Shain W. 1988. Adenosine stimulates cAMP‐mediated taurine release from LRM55 glial cells. J Neurosci Res 20: 176-181. Magnusson KR, Koerner JF, Larson AA, Smullin DH, Skilling SR, et al. 1991. NMDA‐, kainate‐ and quisqualate‐stimulated

Taurine in neurotransmission release of taurine from electrophysiologically monitored rat hippocampal slices. Brain Res 549: 1-8. Malminen O, Kontro P. 1986. Modulation of the GABA‐benzodiazepine receptor complex by taurine in rat brain membranes. Neurochem Res 11: 85-94. Malminen O, Kontro P. 1987. Actions of taurine on the GABA‐benzodiazepine receptor complex solubilized from rat brain. Neurochem Int 11: 113-117. Marnela KM, Kontro P, Pitka¨nen RI, Oja SS. 1980. Free amino acids in synaptic subfractions of bovine brain. Acta Univ Ouluensis [A] 97, Biochem 29: 11-16. Matsuda T, Takuma K, Kishida Y, Azuma J, Baba A. 1996. Protective effect of taurine against reperfusion injury in cultured rat astrocytes. Adv Exp Med Biol 403: 491-497. McBride WJ, Frederickson RC. 1980. Taurine as a possible inhibitory transmitter in the cerebellum. Fed Proc 39: 2701-2705. McCaslin PP, Yu XZ. 1992. Cyanide selectively augments kainate‐ but not NMDA‐induced release of glutamate and taurine. Eur J Pharmacol 228: 73-75. McCool BA, Botting SK. 2000. Characterization of strychnine‐sensitive glycine receptors in acutely isolated adult rat basolateral amygdala neurons. Brain Res 859: 341351. Medina JH, De Robertis E. 1984. Taurine modulation of the benzodiazepine‐g‐aminobutyric acid receptor complex in brain membranes. J Neurochem 42: 1212-1217. Melamed N, Kanner BI. 2004. Transmembrane domains I and II of the g‐aminobutyric acid transporter GAT‐4 contain molecular determinants of substrate specificity. Mol Pharmacol 65: 1452-1461. Mene´ndez N, Solı´s JM, Herreras O, Galarreta M, Conejero C, et al. 1993. Taurine release evoked by NMDA receptor activation is largely dependent on calcium mobilization from intracellular stores. Eur J Neurosci 5: 1273-1279. Miele M, Berners M, Boutelle MG, Kusakabe H, Fillenz M. 1996. The determination of the extracellular concentration of brain glutamate using quantitative microdialysis. Brain Res 707: 131-133. Miyamoto TA, Miyamoto KJ. 1999. Does adenosine release taurine in the A1‐receptor‐rich hippocampus? J Anesth 13: 94-98. Molchanova S, Ko¨o¨bi P, Oja SS, Saransaari P. 2004a. Interstitial concentrations of amino acids during global forebrain ischemia and potassium‐evoked spreading depression. Neurochem Res 29: 1519-1527. Molchanova S, Oja SS, Saransaari P. 2004b. Characteristics of basal taurine release in the rat striatum measured by microdialysis. Amino Acids 27: 261-268. Molchanova SM, Oja SS, Saransaari P. 2005. Mechanisms of enhanced taurine release under Ca2þ depletion. Neurochem Int 47: 343-349.

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Mori M, Ga¨hwiler BH, Gerber U. 2002. b‐Alanine and taurine as endogenous agonists at glycine receptors in rat hippocampus in vitro. J Physiol 539: 191-200. Nadi NS, McBride WJ, Aprison MH. 1977. Distribution of several amino acids in regions of the cerebellum of the rat. J Neurochem 28: 453-455. Nagelhuis EA, Lehmann A, Ottersen OP. 1993. Neuronal‐glial exchange of taurine during hypo‐osmotic stress: A combined immunocytochemical and biochemical analysis in rat cerebellar cortex. Neuroscience 54: 615-631. Napias C, Bergman MO, Van Ness PC, Greenlee DV, Olsen RW. 1980. GABA binding in mammalian brain: Inhibition by endogenous GABA. Life Sci 27: 1001-1011. Neuringer M, Sturman JA, Wen GY, Wisniewski HM. 1985. Dietary taurine is necessary for normal retinal development in monkeys. Prog Clin Biol Res 179: 53-62. O’Byrne MB, Tipton KF. 2000. Taurine‐induced attenuation of MPPþ neurotoxicity in vitro: A possible role for the GABAA subclass of GABA receptors. J Neurochem 74: 2087-2093. Oja SS, Kontro 1978. Neurotransmitter actions of taurine in the central nervous system. Taurine and Neurological Disorders. Barbeau A, Huxtable RJ, editors. New York: Raven Press; pp. 181-200. Oja SS, Kontro P. 1981. Oxidation of hypotaurine in vitro by mouse liver and brain tissues. Biochim Biophys Acta 677: 350-357. Oja SS, Kontro P. 1983. Taurine. Handbook of Neurochemistry, Vol. 3, 2nd edn. Lajtha A, editor. New York: Plenum Press; pp. 501-533. Oja SS, Kontro P. 1989. Release of endogenous taurine and g‐aminobutyric acid from brain slices from the adult and developing mouse. J Neurochem 52: 1018-1024. Oja SS, La¨hdesma¨ki P. 1974. Is taurine an inhibitory neurotransmitter? Med Biol 52: 138-143. Oja SS, Saransaari P. 1992. Cell volume changes and taurine release in cerebral cortical slices. Adv Exp Med Biol 315: 369-374. Oja SS, Saransaari P. 1995. Chloride ions, potassium stimulation and release of endogenous taurine from cerebral cortical slices from 3‐day‐old and 3‐month‐old mice. Neurochem Int 27: 313-318. Oja SS, Saransaari P. 2000. Modulation of taurine release by glutamate receptors and nitric oxide. Prog Neurobiol 62: 407-425. Oja SS, Holopainen I, Kontro P. 1990a. Stimulated taurine release from different brain preparations: Changes during development and aging. Prog Clin Biol Res 351: 277-287. Oja SS, Korpi ER, Saransaari P. 1990b. Modification of chloride flux across brain membranes by inhibitory amino acids in developing and adult mice. Neurochem Res 15: 797-804.

339

340

13

Taurine in neurotransmission

Oja SS, Kontro P, La¨hdesma¨ki P. 1977. Amino acids as inhibitory neurotransmitters. Prog Pharmacol 1/3: 1-119. Okamoto K, Kimura H, Sakai Y. 1983a. Taurine‐induced increase of the Cl
conductance of cerebellar Purkinje cell dendrites in vitro. Brain Res 259: 319-323. Okamoto K, Kimura H, Sakai Y. 1983b. Evidence for taurine as an inhibitory neurotransmitter in cerebellar stellate interneurons: Selective antagonism by TAG (6‐aminomethyl‐4H1,2,4‐benzothiadiazine‐1,1‐dioxide). Brain Res 265: 163-168. Okamoto K, Kimura H, Sakai Y. 1983c. Antagonistic action of 6‐aminomethyl‐4H1,2,4‐benzothiadiazine‐1,1‐dioxide (TAG), and evidence for a transmitter role of taurine in stellate interneurons in the cerebellum. Prog Clin Biol Res 125: 151-160. Olson JE, Martinho E Jr. 2006. Taurine transporter regulation in hippocampal neurons. Adv Exp Med Biol 583: 307-314. O’Neill RD. 1986. Effects of intranigral injection of taurine and GABA on striatal dopamine release monitored voltammetrically in the unanaesthetized rat. Brain Res 382: 28-32. Pasantes‐Morales H, Schousboe A. 1989. Release of taurine from astrocytes during potassium‐evoked swelling. Glia 2: 45-50. Pasantes‐Morales H, Moran J, Schousboe A. 1990. Volume‐ sensitive release of taurine from cultured astrocytes: Properties and mechanism. Glia 4: 427-432. Paula‐Lima AC, De Felice FG, Brito‐Moreira J, Ferreira ST. 2005. Activation of GABAA receptors by taurine and muscimol blocks the neurotoxicity of b‐amyloid in rat hippocampal and cortical neurons. Neuropharmacology 49: 1140-1148. Pierobon P, Minei R, Porcu P, Sogliano C, Tino A, et al. 2001. Putative glycine receptors in Hydra: A biochemical and behavioural study. Eur J Neurosci 14: 1659-1666. Pow DV, Sullivan R, Reye P, Hermanussen S. 2002. Localization of taurine transporters, taurine and 3H taurine accumulation in the rat retina, pituitary, and brain. Glia 37: 153-168. Puopolo M, Kratskin I, Belluzzi O. 1998. Direct inhibitory effect of taurine on relay neurons of the rat olfactory bulb. Neuroreport 9: 2319-2322. Quinn MR, Harris CL. 1995. Taurine allosterically inhibits binding of [35S]‐t‐butyl‐bicyclophosphonothionate (TBPS) to rat brain synaptic membranes. Neuropharmacology 34: 1607-1613. Quinn MR, Miller CI. 1992. Taurine allosterically modulates flunitrazepam binding to synaptic membranes. J Neurosci Res 33: 136-141. Rabe H, Picard R, Uusi‐Oukari M, Hevers W, Lu¨ddens H, et al. 2000. Coupling between agonist and chloride ionophore sites of GABAA receptor: Agonist/antagonist efficacy of 4‐PIOL. Eur J Pharmacol 409: 233-242.

Rajendra S, Lynch JW, Pierce KD, French CR, Barry PH, et al. 1995. Mutation of an arginine residue in the human glycine receptor transforms b‐alanine and taurine from agonists into competitive antagonists. Neuron 14: 169-175. Ramamoorthy D, Del Monte MA, Leibach FH, Ganapathy V. 1994. Molecular identity and calmodulin‐mediated regulation of the taurine transporter in a human retinal pigment epithelial cell line. Curr Eye Res 13: 523-529. Rebel G, Petegnief V, Lleu PL, Gupta RC, Gue´rin P, et al. 1994. New data on the regulation of taurine uptake in cultured nervous cells. Adv Exp Med Biol 359: 225-233. Rogers KL, Philibert RA, Dutton GR. 1991. Kþ‐Stimulated amino acid release from cultured cerebellar neurons: Comparison of static and dynamic stimulation paradigms. Neurochem Res 16: 899-904. Rosati AM, Traversa U, Lucchi R, Poli A. 1995. Biochemical and pharmacological evidence for the presence of A1 but not A2a adenosine receptors in the brain of the low vertebrate teleost Cerassium auratus (goldfish). Neurochem Int 26: 411-423. Ross CD, Godfrey DA, Parli JA. 1995. Amino acid concentrations and selected enzyme activities in rat auditory, olfactory, and visual systems. Neurochem Res 20: 1483-1490. Ruotsalainen M, Ahtee L. 1996. Intrastriatal taurine increases striatal extracellular dopamine in a tetrodotoxin‐sensitive manner in rats. Neurosci Lett 212: 175-178. Ruotsalainen M, Heikkila¨ M, Lillsunde P, Seppa¨la¨ T, Ahtee L. 1996. Taurine infused intrastriatally elevates, but intranigrally decreases striatal extracellular dopamine concentration in anaesthetized rats. J Neural Transm 103: 935-946. Ruotsalainen M, Majasaari M, Salima¨ki J, Ahtee L. 1998. Locally infused taurine, GABA and homotaurine alter differentially the striatal extracellular concentration of dopamine and its metabolites in rats. Amino Acids 15: 117-134. Salima¨ki J, Scriba G, Piepponen TP, Rautolahti N, Ahtee L. 2003. The effects of systemically administered taurine and N‐pivaloyltaurine on striatal extracellular dopamine and taurine in freely moving rats. Naunyn‐Schmiedebergs Arch Pharmacol 368: 134-141. Saransaari P, Oja SS. 1991. Excitatory amino acids evoke taurine release from cerebral cortex slices from adult and developing mice. Neuroscience 45: 451-459. Saransaari P, Oja SS. 1992. Release of GABA and taurine from brain slices. Prog Neurobiol 38: 455-482. Saransaari P, Oja SS. 1993. Phencyclidine binding sites in mouse cerebral cortex during development and ageing: Effects of inhibitory amino acids. Mech Ageing Dev 68: 125-136. Saransaari P, Oja SS. 1994. Taurine release from mouse hippocampal slices: Effects of glutamatergic substances and hypoxia. Adv Med Exp Biol 359: 279-287. Saransaari P, Oja SS. 1997a. Taurine release from the developing and ageing hippocampus: Stimulation by agonists of

Taurine in neurotransmission ionotropic glutamate receptors. Mech Ageing Dev 99: 219-232. Saransaari P, Oja SS. 1997b. Glutamate‐agonist‐evoked taurine release from the adult and developing mouse hippocampus in cell‐damaging conditions. Amino Acids 9: 323-335. Saransaari P, Oja SS. 1998a. Mechanisms of ischemia‐induced taurine release in mouse hippocampal slices. Brain Res 807: 118-124. Saransaari P, Oja SS. 1998b. Release of endogenous glutamate, aspartate, GABA and taurine from hippocampal slices from adult developing mice in cell‐damaging conditions. Neurochem Res 23: 563-570. Saransaari P, Oja SS. 1999a. Taurine release modified by nitric oxide‐generating compounds in the developing and adult mouse hippocampus. Neuroscience 89: 1103-1111. Saransaari P, Oja SS. 1999b. Taurine release is enhanced in cell‐damaging conditions in cultured cerebral cortical astrocytes. Neurochem Res 24: 1523-1529. Saransaari P, Oja SS. 1999c. Involvement of metabotropic glutamate receptors in taurine release in the adult and developing hippocampus. Amino Acids 16: 165-179. Saransaari P, Oja SS. 1999d. Enhanced taurine release in cultured cerebellar granule cells in cell‐damaging conditions. Amino Acids 17: 323-334. Saransaari P, Oja SS. 1999e. Characteristics of ischemia‐ induced taurine release in the developing mouse hippocampus. Neuroscience 94: 949-954. Saransaari P, Oja SS. 2000. Modulation of the ischemia‐ induced taurine release by adenosine receptors in the developing and adult mouse hippocampus. Neuroscience 97: 425-430. Saransaari P, Oja SS. 2002. Taurine release in the developing and adult mouse hippocampus: Involvement of cyclic guanosine monophosphate. Neurochem Res 27: 15-20. Saransaari P, Oja SS. 2003a. Interactions of taurine and adenosine in the mouse hippocampus in normoxia and ischemia. Adv Exp Med Biol 526: 445-451. Saransaari P, Oja SS. 2003b. Characterization of N‐methyl‐D‐ aspartate‐evoked taurine release in the developing and adult mouse hippocampus. Amino Acids 24: 213-221. Saransaari P, Oja SS. 2006. Characteristics of taurine release in slices from adult and developing mouse brain stem. Amino Acids 31: 35-43. Satsu H, Kobayashi Y, Yokohama T, Terasawa E, Shimizu M. 2002. Effect of dietary sulfur amino acids on the taurine content of rat tissues. Amino Acids 23: 447-452. Schaffer SW, Azuma J, Takahashi K, Mozaffari M. 2003. Why is taurine cytoprotective? Adv Exp Med Biol 526: 307-321. Scheller D, Szathmary S, Kolb J, Tegtmeier F. 2000a. Observations on the relationship between the extracellular changes of taurine and glutamate during cortical spreading

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depression, during ischemia, and within the area surrounding a thrombotic infarct. Amino Acids 19: 571-583. Scheller D, Korte M, Szathmary S, Tegtmeier F. 2000b. Cerebral taurine release mechanisms in vivo: Pharmacological investigation in rats using microdialysis for proof of principle. Neurochem Res 25: 801-807. Schmieden V, Betz H. 1995. Pharmacology of the inhibitory glycine receptor: Agonist and antagonist actions of amino acids and piperidine carboxylic acid compounds. Mol Pharmacol 48: 919-927. Schmieden V, Grenningloh G, Schofield PR, Betz H. 1989. Functional expression in Xenopus oocytes of the strychnine binding 48 kDa subunit of the glycine receptor. EMBO J 8: 695-700. Schmieden V, Kuhse J, Betz H. 1992. Agonist pharmacology of neonatal and adult glycine receptor a subunits: Identification of amino acid residues involved in taurine activation. EMBO J 11: 2025-2032. Segovia G, Del Arco A, Mora F. 1997. Endogenous glutamate increases extracellular concentrations of dopamine, GABA, and taurine through NMDA and AMPA/kainate receptors in striatum of the freely moving rat: A microdialysis study. J Neurochem 69: 1476-1483. Seki Y, Feustel PJ, Keller RW Jr, Tranmer BI, Kimelberg HK. 1999. Inhibition of ischemia‐induced glutamate release in rat striatum by dihydrokainate and an anion channel blocker. Stroke 30: 433-440. Simpson RK Jr, Robertson CS, Goodman JC. 1991. Segmental release of amino acid neurotransmitters from transcranial stimulation. Neurochem Res 16: 89-94. Smith KE, Borden LA, Wang CH, Hartig PR, Branchek TA, et al. 1992. Cloning and expression of a high affinity taurine transporter from rat brain. Mol Pharmacol 42: 563-569. Solı´s JM, Herranz AS, Herreras O, Mun˜ozMD, Martı´n del Rı´o R, 1986. Variation of potassium ion concentrations in the rat hippocampus specifically affects extracellular taurine level. Neurosci Lett 66: 263-268. Song Z, Hatton GI. 2003. Taurine and the control of basal hormone release from rat neurohypophysis. Exp Neurol 183: 330-337. Steiner FA. 1971. Neurotransmitter und Neuromodulatoren. Technik und Resultate der Mikroelektrophorese im Nervensystem. Georg Thieme, Stuttgart. Stipanuk MH, Londono M, Lee JI, Hu M, Yu AF. 2002. Enzymes and metabolites of cysteine metabolism in nonhepatic tissues of rats show little response to changes in dietary protein or sulfur amino acid levels. J Nutr 132: 3369-3378. Sturman JA. 1993. Taurine in development. Physiol Rev 73: 119-147. Sturman JA, Moretz RC, French JH, Wisniewski HM. 1985. Taurine deficiency in the developing cat: Persistence of

341

342

13

Taurine in neurotransmission

the cerebellar external granule cell layer. J Neurosci Res 13: 403-416. Sundstro¨m E, Mo LL, Seiger A˚. 1995. In vivo studies on NMDA‐evoked release of amino acids in the rat spinal cord. Neurochem Int 27: 185-193. Sung DY, Walthall WW, Derby CD. 1996. Identification and partial purification of putative taurine receptor proteins from the olfactory organ of the spiny lobster. Comp Biochem Physiol B Biochem Mol Biol 115: 19-26. Sved AF, Curtis JT. 1993. Amino acid neurotransmitters in nucleus tractus solitarius: An in vivo microdialysis study. J Neurochem 61: 2089-2098. Tang XW, Deupree DL, Sun Y, Wu JY. 1996. Biphasic effect of taurine on excitatory amino acid‐induced neurotoxicity. Adv Exp Med Biol 403: 499-505. Tang XW, Yarom M, Carlson RG, Vander Velde D, Huang PY, et al. 1993. Isolation of endogenous modulators for the GABAA and taurine binding. Neurochem Int 23: 485-493. Tappaz ML. 2004. Taurine biosynthetic enzymes and taurine transporter: Molecular identification and regulations. Neurochem Res 29: 83-96. Torp R, Andine´ P, Hagberg H, Karagu¨lle T, Blackstad TW, et al. 1991. Cellular and subcellular redistribution of glutamate‐, glutamine‐ and taurine‐like immunoreactivities during forebrain ischemia: A semiquantitative electron microscopic study in rat hippocampus. Neuroscience 41: 433-447. Trenkner E, El Idrissi A, Dumas R, Rabe A. 1998. Functional consequences of calcium uptake modulation by taurine in vivo and in vitro. Adv Exp Med Biol 442: 277-284. Tuz K, Pen˜a‐Segura C, Franco R, Pasantes‐Morales H. 2004. Depolarization, exocytosis and amino acid release evoked by hyposmolarity from cortical synaptosomes. Eur J Neurosci 19: 916-924. Uchida S, Kwon HM, Yamauchi A, Preston AS, Marumo F, et al. 1992. Molecular cloning of the cDNA for an MDCK cell Naþ‐ and Cl
‐dependent taurine transporter that is regulated by hypertonicity. Proc Natl Acad Sci USA 89: 8230-8234. Vinnakota S, Qian X, Egal H, Sarthy V, Sarkar HK. 1997. Molecular characterization and in situ localization of a mouse retinal taurine transporter. J Neurochem 69: 22382250. Wahl P, Elster L, Schousboe A. 1994. Identification and function of glycine receptors in cultured cerebellar granule cells. J Neurochem 62: 2457-2463. Walberg F, Ottersen OP, Rinvik E. 1990. GABA, glycine, aspartate, glutamate and taurine in the vestibular nuclei: An immunocytochemical investigation in the cat. Exp Brain Res 79: 547-563. Wang DS, Xu TL, Pang ZP, Li JS, Akaike N. 1998. Taurine‐ activated chloride currents in the rat sacral dorsal commissural neurons. Brain Res 792: 41-47.

Wang F, Xiao C, Ye JH. 2005. Taurine activates excitatory non‐ synaptic glycine receptors on dopamine neurons in ventral tegmental area of young rats. J Physiol 565: 503-516. Wu H, Jin Y, Wei J, Jin H, Sha D, et al. 2005. Mode of action of taurine as a neuroprotector. Brain Res 1038: 123-131. Wu JY, Chen W, Tang XW, Jin H, Foos T, et al. 2000. Mode of action of taurine and regulation dynamics of its synthesis in the CNS. Adv Exp Med Biol 483: 35-44. Wu ZY, Xu TL. 2003. Taurine‐evoked chloride current and its potentiation by intracellular Ca2þ in immature rat hippocampal CA1 neurons. Amino Acids 24: 155-162. Yarborough GG, Singh DK, Taylor DA. 1981. Neuropharmacological characterization of a taurine antagonist. J Pharmacol Exp Ther 219: 604-613. Ye GI, Tse ACO, Wh. Yung 1997. Taurine inhibits rat substantia nigra pars reticulate neurons by activation of GABA‐ and glycine‐linked chloride conductance. Brain Res 749: 175-179. Ye JH, Wang F, Krnjevic´ K, Wang W, Xiong ZG, et al. 2004. Presynaptic glycine receptors on GABAergic terminals facilitate discharge of dopaminergic neurons in ventral tegmental area. J Neurosci 24: 8961-8974. Yingcharoen K, Rinvik E, Storm‐Mathisen J, Ottersen OP. 1989. GABA, glycine, glutamate, aspartate and taurine in the perihypoglossal nuclei: An immunocytochemical investigation in the cat with particular reference to the issue of amino acid localization. Exp Brain Res 78: 345-357. Yoshida M, Fukuda S, Tozuka Y, Miyamoto Y, Hisatsune T. 2004. Developmental shift in bidirectional functions of taurine‐sensitive chloride channels during cortical circuit formation in postnatal mouse brain. J Neurobiol 60: 166-175. Yu K, Ge SY, Ruan DY. 2003. Fe2þ decreases the taurine‐ induced Cl
current in acutely dissociated rat hippocampal neurons. Brain Res 960: 25-35. Zhao P, Huang YL, Cheng JS. 1999. Taurine antagonizes calcium overload induced by glutamate or chemical hypoxia in cultured rat hippocampal neurons. Neurosci Lett 268: 25-28. Zheng L, Godfrey DA, Waller HJ, Godfrey TG, Chen K, et al. 2000. Effects of high‐potassium‐induced depolarization on amino acid chemistry of the dorsal cochlear nucleus in rat brain slices. Neurochem Res 25: 823-835. Zhu WJ, Vicini S. 1997. Neurosteroid prolongs GABAA channel deactivation by altering kinetics of desensitized states. J Neurosci 17: 4022-4031. Zielin´ska M, Hilgier W, Borkowska HD, Oja SS, Saransaari P, et al. 2002. Ammonia‐induced extracellular accumulation of taurine in the rat striatum in vivo: Role of ionotropic glutamate receptors. Neurochem Res 27: 37-42. Zielin´ska M, Law RO, Albrecht J. 2003. Excitotoxic mechanism of cell swelling in rat cortical slices treated acutely with ammonia. Neurochem Int 43: 299-303.

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The Endocannabinoid System

B. S. Basavarajappa . R. Yalamanchili . T. B. Cooper . B. L. Hungund

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Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 345

2 2.1 2.1.1 2.1.2 2.1.3 2.1.4 2.1.5 2.1.6 2.1.7 2.2 2.2.1 2.2.2 2.3 2.3.1 2.4 2.5 2.5.1 2.5.2 2.5.3 2.5.4 2.5.5 2.5.6

Endocannabinoid Systems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 346 Cannabinoid Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 346 Glycosylation Sites of Cannabinoid Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 348 Phosphorylation Sites of Cannabinoid Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 349 Cannabinoid Receptor Knockout Mice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 349 Polymorphic Structure of Cannabinoid Receptor Genes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 351 Chromosomal Mapping of the Cannabinoid Receptor Genes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 352 Localization of Cannabinoid Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 352 Signal Transduction Mechanism of Cannabinoid Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 353 Endocannabinoids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 355 Anandamide . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 356 2‐Arachidonylglycerol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 358 Fatty Acid Amide Hydrolase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 361 Localization and Distribution of Fatty Acid Amide Hydrolase in the Brain . . . . . . . . . . . . . . . . . . . 362 Endocannabinoids Uptake . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 363 Roles of Endocannabinoids in Central Nervous System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 364 Regulation of g‐Aminobutyric Acid Transmission . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 364 Regulation of Glutamate Transmission . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 365 Mechanism of Inhibition of Neurotransmitter Release . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 366 Release of Endocannabinoids by Activation of Other Neurotransmitter Receptors . . . . . . . . . . . . 366 Endocannabinoids and Synaptic Plasticity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 367 Role of Endocannabinoid System in Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 367

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Therapeutic Opportunity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 370

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Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 371

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Abstract: The endocannabinoid system is composed of cannabinoid receptors, their endogenous ligands, the endocannabinoids, and the enzymes that produce and inactivate them. Endocannabinoids are a new family of lipid mediators, which includes amides, esters, and ethers of long‐chain polyunsaturated fatty acids. Endocannabinoids engage the cell surface receptors that are targeted by D9‐tetrahydrocannabinol (D9‐THC), the active principle of Cannabissativa preparations like hashish and marijuana. The pathways leading to the synthesis and release of endocannabinoids from neuronal and nonneuronal cells are still rather uncertain. Instead, they are synthesized on demand through cleavage of membrane precursors and are involved in various short‐range signaling processes. In the brain, they combine with CB1 cannabinoid receptors (CB1Rs) on axon terminals to regulate ion channel activity and neurotransmitter release. The endocannabinoid system is shown to be involved in an increasing number of pathological conditions. Their ability to modulate synaptic efficacy has a wide range of functional consequences and provides unique therapeutic possibilities. In this chapter, we aim to provide recent progress in our understanding of the endocannabinoid system in the brain. First, the structure, anatomical distribution, and signal transduction mechanism of cannabinoid receptors are described. The synthetic pathways of endocannabinoids are discussed, along with the putative mechanisms of their release, uptake, and degradation. Finally, the possible role of the endocannabinoid system in the central nervous system is discussed in relation to neurotransmitter release, synaptic plasticity, and the therapeutic role in various disease conditions including alcohol abuse. List of Abbreviations: AA, arachidonic acid; AC, adenylyl cyclase; ACEA, arachidonyl‐20 ‐chloroethylamide/(all Z)‐N‐(2‐cycloethyl)‐5,8,11,14‐eicosatetraenamide; AD, Alzheimer’s disease; A, b‐amyloid; AM251, N‐(piperin‐1‐yl)‐5‐(4‐iodophenyl)‐1‐(2,4‐dichlorophenyl)‐4‐methyl‐1H‐pyrazole‐3‐carboxamide; AM404, N‐(4‐hydroxyphenyl)‐eicosa‐5,8,11,14‐tetraenamide; AMT, anandamide membrane transporter; APE, N‐arachidonoyl phosphatidylethanolamide; BAPTA‐AM, 1,2‐bis‐(2‐aminophenoxy)ethane‐N,N,N’, N’‐tetraacetic acid‐acetoxymethyl ester; cAMP‐Kin, cAMP‐dependent protein kinase; CB1 or CB2, cannabinoid 1 or 2; CBD, cannabidiol; CDTA, calcium‐dependent transacylase; CGA, chorionic gonadotropin; CNS, central nervous system; CP‐55,940, (1R,3R,4R)‐3‐[2‐hydroxy‐4‐(1,1‐dimethylheptyl)phenyl]‐4‐(3‐ hydroxypropyl)cyclohexan‐1‐ol; cPLA2, cytoplasmic phospholipase A2; CPP, conditioned place preference; Cpu, caudate putamen; DA, dopamine; DAG, diacylglycerol; D9‐THC, D9‐tetrahydrocannabinol; DHPG, (S)‐3,5‐dihydroxyphenylglycine; DSI, depolarization‐induced suppression of inhibition; EP, entopeduncular nucleus; EPSC, excitatory postsynaptic currents; FAAH, fatty acid amide hydrolase; FAK, focal adhesion kinase; 5‐HT, 5‐hydroxytryptamine (serotonin); Fyn, src‐family kinases p59; GABA, gamma‐aminobutyric acid; GPCR, G protein‐coupled receptor; GP, globus pallidus; GPe or GPi, external or internal globus pallidus; GPe or GPi, external or internal globus pallidus; GTPS, guanosine 50 ‐O‐(3‐thio)triphosphate; HD, Huntington’s disease; HU‐210, D8‐tetrahydrocannabinol dimethyl heptyl; HD, Huntington’s disease; HU‐ 211, dexanabinol; N-methyl‐D‐aspartate receptor; HU‐211, dexanabinol; IPSC, inhibitory postsynaptic currents; JNK, c‐Jun N‐terminal kinase; LTD, long‐term depression; LTP, long‐term potentiation; MAPK, mitogen activated protein kinase; MetAEA, R‐(þ)‐methanandamide; MGL, monoacylglyceride lipase; mGluRs, metabotropic glutamate receptors; MS, multiple sclerosis; NAc, nucleus accumbens; NADA, N‐arachidonyl‐dopamine; PLD, phospholiapse D; N‐ArPE, N‐arachidonylphosphatidylethanolamine; NMDA receptor, N‐methyl‐D‐aspartate receptor; PD, Parkinson’s disease; PEA, palmitoylethanolamide; PE, phosphatidylethanolamine; PI3K, phosphatidylinositol‐3 kinase; PP2, Src tyrosine kinase inhibitor; PKA, cAMP‐dependent protein kinase; PLA1, phospholipase A1; PLC, Phospholipase C; PL, phospholipase; RFLP, restriction fragment length polymorphism; SNR, substantia nigra pars reticulata; SR141716, N‐(piperidin‐1‐yl)‐5‐(4‐chlorophenyl)‐1‐(2,4‐dichlorophenyl)‐4‐methyl‐1H‐pyrazole‐3‐carboximide hydrochloride (rimonabant); SR144528, N‐((1S)‐endo‐1,3,3‐trimethyl bicyclo heptan‐2‐yl]‐5‐(4chloro‐3‐methylphenyl)‐1‐(4‐methylbenzyl)‐pyrazole‐3‐carboxamide); SSRP, simple sequence repeat polymorphism; TTX, tetrodotoxin; 2‐AG, 2‐arachidonoylglycerol; UCM707, N‐(3‐furanylmethyl)‐5Z,8Z,11Z,14Z‐eicosatetraenamide; URB597, cyclohexyl carbamic acid 30 ‐carbamoyl‐biphenyl‐3‐yl ester; VMN, ventromedial nucleus of the hypothalamus; VTA, ventral tegamental area; VTA, ventral tegmental area; WIN 55,212‐2, R‐(þ)‐[2,3‐dihydro‐5‐methyl‐3‐[(morpholinyl)methyl]pyrrolo‐[1,2,3‐de]‐1,4‐benzoxazinyl]‐(1‐naphthalenyl) methanone mesylate

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Introduction

The ancestors of Cannabis originated in Asia, possibly on the more gentle slopes of the Himalayas or the Altai mountains to the north. The exact origin, obscured by Stone Age trails that cross the continent, is not known. The earliest cultural evidence of Cannabis comes from the oldest known Neolithic culture in China, the Yang‐shao, which appeared along the Yellow River valley about 6,500 years ago (Kabelik et al., 1960). While the Chinese were building their hemp culture, the cotton cultures of India and the linen (flax) cultures of the Mediterranean began to learn of Cannabis through expanding trade and from wandering tribes of Aryans, Mongols, and Scythians who had bordered China since Neolithic times. The Aryans (Indo‐ Persians) brought Cannabis culture to India nearly 4,000 years ago. They worshipped the spirits of plants and animals, and marijuana played an active role in their rituals. In China, with the strong influence of philosophic and moralistic religions, the use of marijuana all but disappeared. However, in India, the Aryan religion grew through oral tradition, until it was recorded in the four Vedas, compiled between 1400 and 1000 B.C. In that tradition, unlike that of the Chinese, marijuana was sacred, and the bhangas spirit was appealed to ‘‘for freedom of distress’’ and as ‘‘a reliever of anxiety’’ (from the Atharva Veda) (Aldrich, 1971). A gift from the gods, according to Indian mythology, the magical Cannabis ‘‘lowered fevers, fostered sleep, relieved dysentery, and cured sundry other ills; it also stimulated appetite, prolonged life, quickened the mind, and improved judgment’’ (Schultes, 1967). The Scythians brought Cannabis to Europe by a northern route where remnants of their campsites, from the Altai Mountains to Germany, date back 2,800 years. Seafaring Europe never smoked marijuana extensively, but hemp fiber became a major crop in the history of almost every European country. Pollen analysis dates the cultivation of Cannabis to 400 B.C. in Norway; 150 A.D. in Sweden, and 400 A.D. in Germany and England (Godwin, 1967), although it is believed that the plant was cultivated in the British Isles several centuries earlier (Frazier, 1974). The Greeks and Romans used hemp for rope and sail but imported the fiber from Sicily and Gaul. Moreover, it has been said ‘‘Caesar invaded Gaul in order to tie up the Roman Empire,’’ an allusion to the Romans’ need for hemp. The Cannabis plant was used in Europe mostly to make cordage and fabric, but first attracted European scientists when Napolean’s troops brought back from Egypt intriguing accounts of its psychotropic activity. An 1848 commentary of the British Pharmacopoeia outlined quite accurately the psychotropic effects of Cannabis and pointed out its merit as an antispamodic and analgesic agent (Christison, 1848). In the 1930s and 1940s, the chemical structure of the first phytocannabinoids had been successfully characterized (Loewe, 1950). However, it was not until 1964 (Gaoni and Mechoulam, 1964) that D9‐ tetrahydrocannabinol (D9‐THC, dronabinol), mainly responsible for the pharmacological effects of the Cannabis plant (Dewey, 1986; Hollister, 1986), was stereochemically defined, and synthesized. The psychoactive properties of D9‐THC were recognized immediately, but the drug’s unique chemical structure offered no hints as to its mechanism of action. The hydrophobic nature of D9‐THC delayed experimentation and suggested that the compound might act by influencing membrane fluidity rather than by binding to specific receptor. The development of new classes of potent and selective D9‐THC analogs led to the pharmacological identification of cannabinoid‐sensitive sites in the brain (Devane et al., 1988). In 1990, the CB1R was molecularly cloned from rat brain (Matsuda et al., 1990), and its immune system counterpart, the CB2 cannabinoid receptor, was identified by sequence homology in 1993 (Munro et al., 1993). These major discoveries not only established the mechanism of action of D9‐THC, led to the development of subtype‐selective agonists and antagonists, but they also initiated a hunt for brain‐derived cannabinoid ligands. The first D9‐THC‐like factor to be isolated was a lipid. It was identified as the amide of arachidonic acid (AA) with ethanolamine, and named anandamide after the Sanskrit word for bliss, ananda (Devane et al., 1992). To date, five endocannabinoids have been identified. These are N‐arachidonylethanolamine (anandamide) (Devane et al., 1992), 2‐arachidonylglycerol (2‐AG) (Mechoulam et al., 1995; Sugiura et al., 1995), 2‐arachidonylglycerol ether (noladin ether) (Hanus et al., 2001), O‐arachidonyl‐ethanolamine (virodhamine) (Porter et al., 2002), and N‐arachidonyldopamine (NADA) (Huang et al., 2002) (> Figure 14-1). Cannabinoid receptors and their endogenous ligands together constitute the endocannabinoid system, which is teleologically millions of years of old and has been found in mammals and many other species (De Petrocellis et al., 1999).

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. Figure 14-1 Molecular structure of endocannabinoids that are known to bind to brain cannabinoid receptors. These endocannabinoids share a polyunsaturated fatty acid moiety (arachidonic acid) and a polar head group consisting of ethanolamine or glycerol

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2.1

Cannabinoid Receptors

The marijuana receptor gene was difficult to clone, but evidence for the existence of the receptor has been demonstrated since the 1980s (Devane et al., 1988; Howlett et al., 1988). It has now been shown that cannabinoids have specific receptors. Much of the progress in cannabinoid research has been achieved in the last decade. So far, two cannabinoid receptor subtypes have been cloned. These are named CB1 and CB2. Evidence for other G‐protein‐coupled cannabinoid receptors (‘‘CB3’’ or ‘‘anandamide receptor’’) in brain and in endothelial tissues is mounting in the literature (Jarai et al., 1999; Wagner et al., 1999; Di Marzo et al., 2000c; Breivogel et al., 2001). However, proof from cloning and expression of a new cannabinoid receptor is yet to come. CB1 and CB2 receptors belong to the large superfamily of receptors that couple to G‐protein‐coupled receptors (GPCR) (for recent review) (Basavarajappa, 2005, 2006). Human CB1 and CB2 receptors share 44% overall amino acid identity (for more details see recent review) (Onaivi et al., 2002). The CB2 receptor shares 81% amino acid identity between rat and mouse, and 81% amino acid identity between rat and human. The CB2 receptor transfected in Chinese hamster ovary cells shows high constitutive activity (Bouaboula et al., 1999a). The CB1R is shown to have a high level of ligand‐independent activation (i.e., constitutive activity) in a variety of cells (Bouaboula et al., 1997; Pan et al., 1998; Meschler et al., 2000; Mato et al., 2002). Nearly 70% exists in the inactive state (R) and 30% exists in the activated state (R*) (Kearn et al., 1999). No correlation was observed between the CB1R concentration and receptor activation in most of the brain regions (Breivogel et al., 1997). The reason for this disparity may be due to the existence of non‐CB1R molecular targets for the endocannabinoids. Although significant progress has been achieved in many aspects of the biology of cannabinoids and our knowledge of cannabinoid genomics and proteomics is increasing, the regulation of cannabinoid receptor

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genes is poorly understood. The cDNA sequences encoding CB1‐ or CB2‐like receptors have been reported in the rat (Matsuda et al., 1993), human (Gerard et al., 1991; Munro et al., 1993), mouse (Chakrabarti et al., 1995; Abood et al., 1997), cow (Wessner and GeneBank submission, 1997), cat (GeneBank submission, 1997) (Gebremedhin et al., 1999), Puffer fish (Yamaguchi et al., 1996), leech (Stefano et al., 1997), and newt (Soderstrom and Johnson, 2000). The CB1R gene structure (> Figure 14-2) is polymorphic, with implications not only for substance abuse but also for other neuropsychiatric disorders. The CB1R gene is intronless and similar in mouse, rat, and human. The CB2 receptor gene is also intronless, at least in its coding region (Onaivi et al., 2002). Both CB1 and CB2 receptors are coded by single‐exon genes. Unlike the CB1R gene, which is highly conserved across the human, rat, and mouse species, the CB2 receptor gene is much more divergent (Griffin et al., 2000). There are two subtypes of CB1R genes (FCB1A and FCB1B) which have been cloned from puffer fish. It was found that these two genes showed high homology to the human CB1 but very low homology to the CB2 receptor gene. The amino acid sequences of FCB1A and FCB2B genes are 66.2% identical, and the homology of each gene to human CB1 is 72.2% and 59%, respectively. Transcripts of both the FCB1A and FCB1B are abundant in the brain (Yamaguchi et al., 1996). There are five splicing mRNA variants (CB1A, CB1B, CB1C, CB1D, and CB1E), identified recently in various regions of human brain (Zhang et al., 2004). Each splicing site was shown to contain canonical GT‐AG splice donor/acceptor motif (> Figure 14-2)

. Figure 14-2 The schematic structure of CB1 receptor gene and variants. The gene is mapped with two finished genomic sequences: AL121835 and AL136096. Although, complete mRNA sequence and exact transcript size are unknown, partial human mRNA sequences are available. The known sequences consist of four exons; one of them includes the complete coding region of the gene (exon 4). Three novel exons that were termed as exon 1, 2, and 3 located 50 to the previously described main CB1 receptor gene coding exon (exon 4). The sizes of these exons were given in the parenthesis. These exons were separated by introns I (1.4 kb), II (12.2 kb), and III (2.3 kb). There were five variants of CB1 receptor genes, which were identified so far. For details, see > Section 2.1. [The original figure was modified and reproduced with permission from Emmanuel S Onaivi (Onaivi et al., 2002; Zhang et al., 2004)]

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(Zhang et al., 2004). Two more variants of CB1R (hCB1a and hCB1b) were identified in human tissue. None of the known endogenous ligands of the CB1R identified to date has any significant agonist activity at either of the splice variants (hCB1a and hCB1b) (Ryberg et al., 2005). It is possible that additional endocannabinoids remain unidentified that may have activity at these variant receptors (Ryberg et al., 2005). Like other GPCRs, the primary structures of the cannabinoid receptors are characterized by the seven hydrophobic stretches of 20–25 amino acids predicted to form transmembrane a‐helices, which are connected by alternating extracellular and intracellular loops. It has been reported that the human and rat N‐terminal 28 amino acids in the CB1Rs were similar in the total number of nonpolar, polar, acidic, and basic amino acids. The mouse N‐terminal 28 amino acids differed from the rat and human CB1Rs in number and composition of the total nonpolar and polar amino acids. There are also significant differences in the total nonpolar, polar, acidic, and basic amino acid composition of the N‐terminal 28 amino acids between human CB1 and CB2 receptors. Further, the molecular weights of human, rat, and mouse CB1Rs are similar. Therefore, the amino acid composition of the mammalian CB1Rs shows strong conservation in contrast to molecular weights and amino acid composition of CB2 receptors (Onaivi et al., 2002). In CB1 and CB2 receptors, no cysteines are found within the second extracellular domain, but the third extracellular domain contains two or more cysteines. Cysteine residues appear to stabilize the tertiary structure of the receptor because of their involvement in intramolecular disulfide bridges. In most GPCRs, these cysteines occur in the extracellular domains that lie between two and three in hydrophobic domains, and four and five in hydrophobic domains (second and third extracellular domains, on the assumption that the N‐terminal domain is also extracellular). One more deviation from most other GPCRs is that CB1 and CB2 receptors lack a highly conserved proline residue in the fifth hydrophobic domain (Matsuda, 1997). The structural features of these proteins, which are critical for ligand binding and functional properties, have been evaluated in in vivo and in vitro models (Akinshola et al., 1999a, b).

2.1.1 Glycosylation Sites of Cannabinoid Receptors The transmembrane helix bundle arrangement obtained for the CB1R is similar to that obtained for other GPCRs. Most, but not all, GPCRs are glycoproteins. The consensus sites for N‐glycosylation are mainly concentrated at the N‐terminus of the protein. There are three potential N‐glycosylation sites highly conserved in human, rat, and mouse (> Figure 14-3). The rodent CB1R protein has an additional . Figure 14-3 Potential N‐glycosylation (N‐glycos) and protein kinase sites in human CB1 receptor proteins (PKC, protein kinase C; cAMP‐Kin, camp‐dependent protein kinase; Ca‐kin II, Ca‐dependent protein kinase II). [Reproduced with permission from Emmanuel S Onaivi (Onaivi et al., 2002)]

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potential N‐glycosylation site at the C‐terminal segment that is absent in the human CB1R protein. One potential N‐glycosylation site is present in human and rat CB1R protein, but that site is missing in mouse CB1R (Onaivi et al., 1996). Whether all of these potential N‐glycosylation sites are naturally glycosylated in CB1R proteins is not known. Whether these N‐glycosylation sites are essential for CB1R function has yet to be determined. The human CB1 and CB2 receptors, with some similarities and some differences in their receptor function, appear to differ in the number and distribution of their potential N‐glycosylation sites. In the N‐terminal region, the CB2 receptor has only one potential N‐glycosylation site, whereas the CB1R has five. There is no potential N‐glycosylation site at the C‐terminal segment of the CB2 receptor (Onaivi et al., 1996). The biological significance (if any) of these differences is yet to be determined.

2.1.2 Phosphorylation Sites of Cannabinoid Receptors There are four clusters of potential cAMP‐dependent protein kinase and Ca2þ‐calmodulin‐dependent protein kinase sites in CB1Rs. These clusters are conserved across human, rat, and mouse receptor proteins. There is a single potential protein kinase C site that is also conserved in these CB1Rs (> Figure 14-3). There is no such site in CB2 receptors. The potential N‐terminal cAMP cluster present in CB1R appears to be conserved, and the CB2 receptor has two such potential sites. None of the CB1 and CB2 receptors has any potential protein kinase sites at the C‐terminal regions. The biological significance of these potential protein phosphorylation sites in these receptor molecules is yet to be determined.

2.1.3 Cannabinoid Receptor Knockout Mice CB1 and CB2 receptor knockout mice have been generated. The CB2 receptor knockout mouse was generated (Buckley et al., 1997) to study the function of these receptors in immune cells and immunomodulation. The mice deficient in the CB2 receptor gene were generally healthy and fertile, they cared for their offspring, and in situ hybridization histochemistry demonstrated the absence of CB2 mRNA in the knockout mice (Buckley et al., 1997, 2000). Binding studies on intact spleens and splenic membranes using the highly specific ligand [3H] CP55940 showed significant binding in spleens derived from wild‐type mice but no such binding in spleens from the CB2 receptor mutant mice (Buckley et al., 1997, 2000). It was reported that D9‐THC inhibits helper T‐cell activation by macrophages in wild type, but not in CB2 mutant mice, indicating that cannabinoids inhibit macrophage costimulatory activity and T‐cell activation via the CB2 receptors (Buckley et al., 1997). While these studies continue, mice deficient in the CB2 receptor gene demonstrated that the CB2 receptor is involved in cannabinoid‐induced immunomodulation and not involved in the central nervous system (CNS) effects of cannabinoids (Buckley et al., 2000). These investigators suggested that it might be possible in the future to separate central and peripheral effects of cannabinoids with selective agents acting at the CB2 receptor sites that may be devoid of the psychoactive effects known to be mediated via CB1R activity in the CNS. Two groups have generated CB1R gene knockout mice independently. These two groups produced mutant mice with disrupted CB1R genes by standard homologous recombination techniques similar to that used in the production of the CB2 mutant mice. The first CB1 knockout mice were generated on a CD1 background (Ledent et al., 1999) and showed that the spontaneous locomotor activity in these mutant mice was increased and that they did not respond to cannabinoid drugs, suggesting that the CB1R was responsible for mediating the analgesic, reinforcement, hypothermic, hypolocomotive, and hypotensive effects of cannabinoids. It was also shown that the acute effects of opiates were unaffected but that the reinforcing properties of morphine and the severity of the withdrawal syndrome were strongly reduced in these knockout mice. The second knockout mouse was generated on a C57BL background (Steiner et al., 1999; Zimmer et al., 1999) and showed that the CB1 mutant mice appear healthy and fertile but had significantly higher mortality rates and showed reduced spontaneous locomotor activity, increased immobility, and hypoanalgesia when compared to the wild‐type littermates (Steiner et al., 1999; Zimmer et al., 1999). In these CB1

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mutant mice, D9‐THC‐induced catalepsy, hypomobility, and hypothermia were absent, but D9‐THC‐ induced analgesia in the tail‐flick test and other behavioral (licking of the abdomen) and physiological (diarrhea) responses to THC were still present. Thus, there were behavioral similarities and differences among CB1 knockout mice on CD1 and C57BL background. It appears that the differences in responses by the mutant mice from the two groups might be related to methodological differences derived from using different laboratory techniques. The groups, however, differed in their findings on the baseline motility of the CB1 mutants. It was found that the CB1 mutant mice on CD1 background exhibited higher levels of spontaneous locomotion, even when placed in fear‐inducing novel environments (such as elevated plus‐ maze and open field). In contrast, the CB1 mutant mice on C57BL background displayed reduced activity in the open‐field test and an increased tendency to be cataleptic. In the basal ganglia, a brain structure with high levels of CB1R that is important for sensorimotor and motivational aspects of behavior was shown to display significantly increased levels of substance P, dynorphin, enkephalin, and GAD67 gene product expression that may account for the alterations in spontaneous activity observed in the CB1 mutant mice. Overall, these findings provide many valuable insights into cannabinoid mechanisms, despite some differences in reports on behavior in these CB1R gene knockout mice. There is, therefore, general agreement that the CB1R plays a key role in mediating most, but not all, CNS effects of cannabinoids. The biological consequences of inactivating the CB1 and/or CB2 receptor genes have continued to be studied intensively. The availability of the cannabinoid knockout mice provides an excellent opportunity to study the biological roles of these genes. CB1R knockout mice exhibited 50% larger long‐term potentiation (LTP) than wild‐type controls, with other properties of the Schaffer collateral‐CA1 synapses (hippocampal model for synaptic changes that are believed to underlie memory at the cellular level), such as paired‐pulse facilitation, remaining unchanged (Bohme et al., 2000). These results suggested that disrupting the CB1R‐ mediated neurotransmission at the genome level produces mutant mice with an enhanced capacity to strengthen synaptic connections in a brain region crucial for memory formation. CB1 knockout mice were able to retain memory in a two‐trial object recognition test for at least 48 h after the first trial, whereas the wild‐type controls lose their capacity to retain memory after 24 h. These data along with previous findings of other investigators suggest that the endogenous cannabinoid systems play a crucial role in the process of memory storage and retrieval (Reibaud et al., 1999; Bohme et al., 2000). This finding is supported by previous data indicating enhanced LTP in mice lacking the CB1R gene (Bohme et al., 2000). These rapid advances in cannabinoid research have continued to add to our knowledge about the biology of marijuana (cannabinoids) in vertebrate and invertebrate systems. These knockout mice lacking cannabinoid receptor genes have also enabled scientists to investigate the interaction of cannabinoids with other neurochemical networks. It was demonstrated that the absence of the CB1R did not modify the antinociceptive effects induced by mu, delta, and kappa opioid agonists, but these mice exhibited a reduction in stress‐induced analgesia (Valverde et al., 2001). These results therefore suggest that the CB1Rs are not involved in the antinociceptive responses to exogenous opioids but that a physiological interaction between the opioid and cannabinoid systems is necessary to allow the development of opioid‐ mediated responses to stress. Further, in a different study, it was shown that morphine did not modify dopamine release in the nucleus accumbens (NAc) of CB1R knockout mice under conditions where it dose‐ dependently stimulates the release of dopamine in the corresponding wild‐type mice (Mascia et al., 1999), indicating that the CB1R regulates mesolimbic dopaminergic transmission in brain areas known to be involved in the reinforcing effects of morphine (Mascia et al., 1999). Recent results provide unequivocal evidence that the acute alcohol‐induced dopamine release in the NAc is in fact mediated by CB1Rs (Hungund et al., 2003). Further, SR141716A blocked alcohol‐evoked dopamine release in the shell of the NAc following alcohol administration (Cohen et al., 2002). The acute alcohol‐induced increase in dopamine in NAc dialysates in C57BL/6 mice was completely inhibited by pretreatment with SR141716A or deletion of CB1Rs in mice (CB1 receptor knockout) (Hungund et al., 2003). These data indicate that the CB1R regulates mesolimbic dopaminergic transmission in brain areas known to be involved in the reinforcing effects of drugs of abuse including alcohol. The genetic homologies between the CB1R and the D2 dopamine receptor (Fritzsche, 2000) and the presynaptic regulation of neurotransmitter release support a closer investigation of the role of endocannabinoids in the physiological control systems of addiction and other neuropyschiatric disorders.

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Other researchers have begun to look at the CB1R knockout mice as potential animal models for schizophrenia (Fritzsche, 2000) and as models for other neuropsychiatric conditions because of the ubiquity, diverse functions, and numerous signal transductions involved in the actions of cannabinoids. Rapid advances in the designing of genetically engineered laboratory animals are producing models that are more effective research tools. As a result, the need for precise genetic characterization of laboratory animals is of primary concern. However, the use of transgenic mouse models for the overexpression of CB1 or CB2 receptor genes (as opposed to CB1 or CB2 receptor gene inactivation or knockout as described above) to study the regulation and site‐specific mechanisms of action of cannabinoids are currently unexplored. For example, the use of cannabinoid transgenics, in which genomic regulatory sequences of interest are coupled to a reporter gene can be used to probe further the mechanism of regulation of the CB1 or CB2 receptor genes. The current lack of information about the promoters, 50 ‐ and 30 ‐UTRs, and other regulatory elements of CB1 or CB2 receptor genes hampers any effort to make such CB1 or CB2 receptor gene construct modification to generate such rodent CB1 or CB2 receptor transgenic models. Obviously, the use of CB1 or CB2 receptor transgenic animals will provide new in vivo systems for studying genetic regulation, development, normal physiology, and apparent dysfunctions associated with the cannabinoid system.

2.1.4 Polymorphic Structure of Cannabinoid Receptor Genes Improved information about cannabinoid receptors and their allelic variants in humans and rodents can add to our understanding of vulnerability to addiction and other neuropsychiatric disorders. Little information is available, however, at the molecular level about cannabinoid receptor gene polymorphisms. Different human cannabinoid receptor gene polymorphisms have been reported. A silent mutation resulting in the substitution from G to A at nucleotide position 1359 in codon 453 (Thr) turned out to be a common polymorphism in the German GTS patient population (Gadzicki et al., 1999). No significant differences were found in allelic distribution between GTS patients and controls within the coding region of the CB1R gene (Gadzicki et al., 1999). The frequencies of this polymorphism are significantly different between Caucasian, African American, and Japanese populations (Onaivi et al., 2002). A HindIII restriction fragment length polymorphism (RFLP) is located in an intron approximately 14 kb in 50 region of the initiation codon of the CB1R gene. A polymorphism consisting of two alleles with 0.23 and 0.77 frequencies in unrelated Caucasians was reported using hybridization of human DNA digested with HindIII (Caenazzo et al., 1991). Another polymorphism associated with the CB1R gene with mental illness and drug abuse in different populations is a simple sequence repeat polymorphism (SSRP) consisting of nine alleles containing (AAT) 12–20 repeat sequences was reported (Dawson, 1995). There was no significant association of the CB1R gene AAT repeat polymorphism with Chinese heroin addicts and there was no evidence that the CB1R gene AAT repeat polymorphism confers susceptibility to heroin abuse with heroin abuse in a Chinese population (Li et al., 2000). The studies to test for linkage with susceptibility to alcohol or drug dependence using the CB1R gene triplet repeat marker suggested a significant association of the CB1R gene with a number of different types of drug dependence (cocaine, amphetamine, cannabis) and intravenous drug use. There was no significant association of this marker with alcohol abuse/dependence in nonHispanic Caucasians (Comings et al., 1997). In addition, there was also a significant association of the triplet repeat marker in the CB1R gene alleles with the P300 event‐related potential that has been implicated in substance abuse (Johnson et al., 1997). The CB1R gene AAT repeat marker was used to test for linkage with schizophrenia, and results indicated no linkage or association of the marker with schizophrenia, indicating that the CB1R gene is not a gene of major etiological effect for schizophrenia (Comings et al., 1997). The CB1R gene is located in human chromosome 6q14‐q15, and it is interesting that previous reports showed evidence for possible linkage to schizophrenia with chromosome 6q markers (Martinez et al., 1999). In addition, evidence exists for a schizophrenia susceptibility locus on chromosome 6q (Cao et al., 1997). Although there were no linkages or associations of the CB1R triplet marker with schizophrenia, it remains to be determined if linkage and association to schizophrenia might exist with other as yet unknown polymorphisms that exist in the CB1R gene structure.

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Three other variants have been reported in the CB1R gene obtained by PCR assay with cDNA from hippocampal tissue taken from patients undergoing neurosurgery for intractable epilepsy (Kathmann et al., 2000). The updated picture (> Figure 14-2) of the CB1/Cnr1 presented in a recent study provides a gene that displays at least four exons that are expressed in hippocampus and other brain regions, spans 25 kb, and produces several transcripts. The gene has novel SNPs, a T insertion/deletion in its exon 3 that encodes 50 untranslated region (UTR) sequences, and a 3813G>A polymorphism in exon 4 sequences that encodes 30 UTR sequences. Characterization of this locus now locates the previously described (AAT)n polymorphism more than 12 kb from the main CB1R amino acid coding exon 4. These data, and the associated data reported here, provide an improved basis to understand the nature of the human CB1R locus and possible roles for its functional variants in human addiction.

2.1.5 Chromosomal Mapping of the Cannabinoid Receptor Genes The genomic location of the human cannabinoid receptor gene was done using genetic linkage mapping, and chromosomal in situ hybridization suggested the presence of the cannabinoid receptor gene on chromosome 6q14‐q15 (Hoehe et al., 1991). This location is very near to the gene encoding the alpha subunit of chorionic gonadotropin (CGA). The mouse CB1 and CB2 receptor genes are located in the proximal arm of chromosome 4 (Stubbs et al., 1996; Onaivi et al., 2002). This location is within a region where other homologs of human 6q genes are located. The genes encoding the peripheral CB2 receptors and a‐L‐fucosidase have been shown to be located near a newly identified common virus integration site, Ev11 (Valk et al., 1997). The site Ev11 is located at the distal end of mouse chromosome 4 in a region that is synthenic with human 1p36, which is in agreement with the report of others that the mouse CB2 receptor gene is also located at the distal end of mouse chromosome 4 (Onaivi et al., 1998). The location of the rat CB1R genes in the rat genome has not been determined but may be expected to fit the rodent–human homology because the CB1R genes are highly conserved in the mammalian species. The physical and genetic localization of the bovine CB1R genes has been mapped to chromosome 9q22 (Pfister‐Genskow et al., 1997). As the neurobiological effects of marijuana and other cannabinoids suggest the involvement of the cannabinoid receptor genes in mental and neurological disturbances, the mapping of the genes will undoubtedly enhance our understanding of the linkage and genetic localization of possible cannabinoid abnormalities.

2.1.6 Localization of Cannabinoid Receptors The CB1R is mainly expressed in brain and spinal cord and thus is often referred to as the ‘‘brain cannabinoid receptor.’’ CB1Rs are among the most abundant GPCRs in brain, their densities being similar to levels of g‐aminobutyric acid (GABA) and glutamate‐gated ion channels. The distribution of cannabinoid receptors within the CNS was first described in a landmark study using quantitative in vitro receptor autoradiography with the radioligand [3H]CP55940 (Herkenham et al., 1991). The distribution of CB1Rs is highly heterogeneous, with the highest densities of receptors present in the outflow nuclei of the basal ganglia, substantia nigra pars reticulata (SNR), and the internal and external segments of the globus pallidus (GP). In addition, very high levels of binding are present in the hippocampus, particularly within the dentate gyrus, and in the molecular layer of the cerebellum. In contrast, there are few CB1Rs in the brain stem. There is a similar distribution of CB1Rs in humans (Glass et al., 1997; Biegon and Kerman, 2001). The highest densities are found in association with limbic cortices, with much lower levels within primary sensory and motor regions, suggesting an important role in motivational (limbic) and cognitive (association) information processing. Recent studies using immunohistochemical approaches with antibodies to either the C‐ (Egertova and Elphick, 2000) or N‐terminal (Pettit et al., 1998; Tsou et al., 1998a) of the CB1R corroborate the earlier autoradiographic studies and provide insight into the subcellular distribution of CB1Rs in areas such as hippocampus and amygdala (Katona et al., 1999, 2001). Combined with electron microscopy and electrophysiology studies, CB1Rs have been shown to be localized presynaptically on GABAergic interneurons (Katona et al., 1999, 2001). This would be consistent with the proposed role of endocannabinoid compounds in modulating neurotransmission. Hence, the anatomy of CB1Rs can provide clues to their function.

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The CB2 receptor is at times referred to as the ‘‘peripheral cannabinoid receptor’’ because of its largely peripheral expression in immune cells. CB2 is not expressed in even moderate abundance in any brain region but is expressed in peripheral tissues including white blood cells (Munro et al., 1993; Facci et al., 1995). CB2 receptor mRNA has been found in spleen, tonsils, and thymus, which are the major tissues of immune cell production and regulation (see Cabral and Dove Pettit, 1998; Howlett et al., 2002 for review). CB2 mRNA has been localized to B and T lymphocytes, natural killer cells, monocytes, macrophages, microglial cells, mast cells, and also cultured cell models of these immune cells. CB2 agonists are generally suppressive of function of these cells, but both CB1 and CB2 receptors might contribute to these effects (Cabral and Dove Pettit, 1998).

2.1.7 Signal Transduction Mechanism of Cannabinoid Receptors Significant and complex information has been accumulated about the associated signal transduction pathway involved with the actions of marijuana and cannabinoids (> Figure 14-4). However, in the context of classical ideas of synaptic signaling, it is important to establish whether CB1R signaling involves postsynaptic

. Figure 14-4 The mechanism of anandamide formation. Stimulation of adenylate cyclase and cAMP‐dependent protein kinase potentiates the N‐acyltransferase (Ca2þ‐dependent transacylase, CDTA). A fatty arachidonic acid chain is transferred by CDTA from the sn‐1 position of phospholipids to the primary amine of phosphatidylethanolamine, in a Ca2þ‐dependent manner, forming an N‐arachidonylphosphatidylethanolamine (N‐arachidonylPE). This N‐arachidonylPE intermediate is then hydrolyzed by a phospholipase D (PLD)‐like enzyme to yield the anandamide (AEA). Once synthesized, AEA can be transported to the outside of the cell through a process that is not yet been characterized. AEA is taken up from the extracellular space, by a transport process that can be blocked by AM404, and is inactivated by fatty acid amide hydrolase (FAAH). AMT, anandamide membrane transporter. For details, see > Section 2.2.1. [Reproduced with permission from Balapal Basavarajappa (2006)]

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receptor or presynaptic receptor or both. Recent electrophysiological and immunocytochemical studies suggest the existence of the CB1R at presynaptic terminals. Immunocytochemical studies showing the loss of receptors in the striatum, pallidum, and nigra following destruction of striatal neurons with ibotenic acid have shown that the CB1R is targeted to the axons and terminals of neurons. Both CB1 and CB2 cannabinoid receptors are coupled with Gi or Go protein, negatively to adenylate cyclase. Adenylate cyclase activity was inhibited by N‐arachidonylethanolamine (AEA), R(þ)‐methanandamide (MetAEA), and 2‐AG in N18TG2 cells (Childers et al., 1994; Pinto et al., 1994; Howlett and Mukhopadhyay, 2000). Upregulation of adenylate cyclase activity was reported without Gi/o coupling (pertussis toxin sensitive) probably through activation of Gs proteins (Glass and Felder, 1997). In another study, the expression of specific isoforms of adenylate cyclase such as I, III, V, VI, or VIII with coexpression of CB1 or CB2 is associated with the inhibition of cyclic AMP accumulation. However, the expression of adenylate cyclase isoforms II, IV, or VII with coexpression of the CB1 or CB2 receptor is associated with stimulation of cAMP accumulation (Rhee et al., 1998). The questions whether CB1R coupling to Gs proteins has physiological importance and whether such coupling increases after Gi or Go protein sequestration by colocalized noncannabinoid Gi/o‐protein‐coupled receptors have yet to be resolved. CB1R coupling to the G‐protein signal transduction pathways in presynaptic nerve terminals transduces the cannabinoid inhibition of adenylate cyclase, by attenuating the production of cyclic AMP. CB1R‐ and Gi/o‐protein‐mediated activation of p42/p44 MAPK was shown in U373MG astrocytoma cells and in host cells expressing recombinant CB1Rs (Bouaboula et al., 1995b). Activation of CB1Rs by D9‐THC and HU‐210 activated p42/p44 MAPK via Gi/o proteins in C6 glioma and primary astrocytes cultures (Sanchez et al., 1998; Guzman and Sanchez, 1999). In WI‐38 fibroblasts, AEA promoted Tyr‐ phosphorylation of extracellular signal‐regulated kinase 2 (ERK2 or p44) and increased MAPK activity mediated through Gi/o proteins (Wartmann et al., 1995). In some cells, CB1R‐mediated activation of MAPK was blocked by the PI3 kinase inhibitor Wartmann, implicating the PI3 kinase in this pathway (Wartmann et al., 1995; Bouaboula et al., 1995b). Endocannabinoids (AEA and 2‐AG) and D9‐THC‐stimulated tyrosine phosphorylation of FAKþ6,7, a neuronal splice isoform of FAK, on several residues including Tyr‐397 in hippocampal slices (Derkinderen et al., 1996; Derkinderen et al., 2001). Cannabinoids increased phosphorylation of p130‐Cas, a protein associated with FAK, but had no effect on PYK2, a tyrosine kinase related to FAK and enriched in hippocampus. Endocannabinoids increased the association of Fyn, but not Src, with FAKþ6,7. These effects were sensitive to manipulation of cAMP‐dependent protein kinase, suggesting that they were mediated by inhibition of a cAMP pathway. PP2, an Src family kinase inhibitor, prevented the effects of cannabinoids on p130‐Cas and on FAKþ6,7 tyrosines 577 and 925, but not 397. These observations suggest that FAK autophosphorylation was upstream of Src family kinases in response to CB1R stimulation (Derkinderen et al., 2001). D9‐THC promoted phosphorylation of Raf‐1 and subsequent translocation to membrane in cortical astrocytes (Sanchez et al., 1998). These observations suggest a pathway whereby CB1R‐mediated release of bg subunits leads to activation of PI3K, resulting in tyrosine phosphorylation and activation of Raf‐1, and the resulting MAPK phosphorylation. Activation of p38 MAPK was observed in CHO cells expressing recombinant CB1Rs (Rueda et al., 2000) and in human vascular endothelial cells having endogenous CB1Rs (Liu et al., 2000). D9‐THC was shown to induce activation of c‐jun N‐terminal kinase (JNK1 and JNK2) in CHO cells expressing recombinant CB1Rs (Rueda et al., 2000). The pathway for JNK activation involves Gi/o protein, PI3K, and Ras (Rueda et al., 2000). MAPK, which is activated by stimulation of CB1Rs is shown to activate the Naþ/Hþ exchanger in CHO cells stably expressing the CB1R (Bouaboula et al., 1999b). AEA‐stimulated activation of MAPK activity was shown to phosphorylate cytoplasmic phospholipase A2 (cPLA2), release of arachidonic acid, and resulting synthesis of prostaglandin E2 in WI‐38 cells (Wartmann et al., 1995). MAPK activation by cannabinoids was shown to induce immediate early gene expression (Krox‐24) in U373MG human astrocytoma cells (Bouaboula et al., 1995a). The suppression of prolactin receptor and trk nerve growth factor receptor synthesis by AEA was shown to be associated with CB1R‐mediated decrease in protein kinase A and increase in MAPK activities (De Petrocellis et al., 1998). CB1R agonists induce the expression of c‐fos and c‐jun in brain (Mailleux et al., 1994; McGregor et al., 1998; Arnold et al., 2001), whether this is mediated by CB1R‐ activated MAPK is not known. D9‐THC‐induced phosphorylation of the transcription factor Elk‐1 is

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mediated by MAPK/ERK (Valjent et al., 2001). D9‐THC and HU‐210 increase glucose metabolism and glycogen synthesis in C6 glioma and astrocytes cultures (Guzman and Sanchez, 1999). The activation of protein kinase B/Akt (isoforms IB) by cannabinoid agonists is mediated by Gi/o and PI3K in U373MG astrocytoma and CHO cells expressing recombinant CB1Rs (Gomez del Pulgar et al., 2000). In addition, CB1R activation has been found to lead to the generation of ceramide (Guzman et al., 2001). This widespread lipid second messenger is known to play an important role in the control of cell fate in the CNS. Studies showed that cannabinoid‐dependent ceramide generation occurs by a G‐protein‐independent process and involves two different metabolic pathways: sphingomyelin hydrolysis and ceramide synthesis de novo. Ceramide in turn mediates cannabinoid‐induced apoptosis, as shown by in vitro and in vivo studies. Thus, the CB1Rs of astrocytes is shown to be coupled to sphingomyelin hydrolysis through the adapter protein (FAN) factor associated with neutral sphingomyelinase activation (Sanchez et al., 2001). L‐type Ca2þ channels are inhibited by CB1R agonist in cat brain arterial smooth muscle cells, which express mRNA and CB1Rs (Gebremedhin et al., 1999). This is blocked by pertussis toxin and SR141716A (Gebremedhin et al., 1999). CB1Rs are also coupled to ion channels through Gi/o proteins, positively to A‐ type and inwardly rectifying potassium channels and negatively to N‐type and P/Q‐type calcium channels and to D‐type potassium channels (Howlett and Mukhopadhyay, 2000; Howlett et al., 2002). The coupling to A‐type and D‐type potassium channels is thought to be through adenylate cyclase (Mu et al., 1999). These are also stimulated by the inhibition of adenylate cyclase by cannabinoids. Because of the decrease in cAMP accumulation, cAMP‐dependent protein kinase (PKA) is inhibited by CB1R activation. Without cannabinoids, PKA phosphorylates the potassium channel protein, by exerting decreased outward potassium current. In the presence of cannabinoids, however, the phosphorylation of the channel by PKA is reduced, which leads to an increased outward potassium current (Childers and Deadwyler, 1996). In addition, cannabinoids can close sodium channels, but whether this effect is receptor‐mediated has yet to be proved. Based on these findings, it has been suggested that cannabinoids play a role in regulation of neurotransmitter release. There is also evidence from experiments with rat hippocampal CA1 pyramidal neurons that CB1Rs are negatively coupled to M‐type potassium channels (Schweitzer, 2000). CB1Rs may also mobilize arachidonic acid and close 5HT3 receptor ion channels (Pertwee, 1997), in certain conditions couple to Gs proteins to activate adenylate cyclase (Calandra et al., 1999) to reduce outward potassium K current, possibly through arachidonic acid‐mediated stimulation of protein kinase C (Hampson et al., 2000). CB1Rs have also been reported to be positively coupled to phospholipase C (PLC) through G proteins in COS‐7 cells cotransfected with CB1Rs and Ga subunits (Ho et al., 1999), also negatively coupled to voltage‐gated L‐type calcium channels in cat cerebral arterial smooth muscle cells (Gebremedhin et al., 1999). CB1Rs on cultured cerebellar granule neurons can work through a PLC‐sensitive mechanism to increase N‐methyl‐D‐aspartate (NMDA)‐elicited calcium release from inositol 1,4,5‐triphosphate‐gated intracellular stores (Netzeband et al., 1999). Activation of CB1Rs by cannabinoid agonists evokes a rapid, transient increase in intracellular free Ca2þ in N18TG2 and NG108‐15 cells (Sugiura et al., 1996c; Sugiura et al., 1997; Sugiura et al., 1999). The CB2 receptor is also coupled to a G protein and negatively coupled to adenylate cyclase.

2.2

Endocannabinoids

As the first plant‐derived cannabinoid, THC, was structurally defined and synthesized over 30 years ago, analogs have been synthesized exogenously, including synthetic cannabinoids like CP55940, HU‐210, and WIN55212 (an aminoalkylindole). All of these bind to receptors (CB1 and CB2) and induce cannabimimetic activity (Howlett et al., 2002). There had been considerable speculation as to whether endogenous ligands existed, especially as CB1 is the most abundantly expressed of neuronal receptors. The identification of endogenous cannabinoid receptors suggested that the brain produces its own chemicals that interact with the CB1Rs during normal brain function, thus suggesting the existence of an endogenous cannabinoid signaling pathway in the brain. Beginning in 1992, two endogenous ligands for mammalian cannabinoid receptors were discovered and characterized. These are AEA and 2‐AG (> Figure 14-1) (Devane et al., 1992; Mechoulam et al., 1995;

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Sugiura et al., 1995). The third ether‐type endocannabinoid, 2‐arachidonylglycerol ether (noladin ether) (> Figure 14-1), has been isolated from CNS and shown to display pharmacological properties similar to AEA (Hanus et al., 2001). The fourth type of endocannabinoid, virodhamine (> Figure 14-1), in contrast to previously described endocannabinoids, is a partial agonist with in vivo antagonist activity at the CB1Rs (Porter et al., 2002). The fifth type of endocannabinoid, NADA, not only binds to CB1Rs but also stimulates vanilloid receptors (VR1) (Huang et al., 2002). Previously, the existence of AEA analogs in chocolate had been demonstrated (di Tomaso et al., 1996). It is thought that chocolate and cocoa contain N‐acylethanolamines (NAE), which are chemically and pharmacologically related to AEA. These lipids could mimic cannabinoid ligands either directly by activating cannabinoid receptors or indirectly by increasing AEA levels (Bruinsma and Taren, 1999). These observations indicate that endocannabinoid analogs exist in plants and animals, and further show the evolutionary conservation of the cannabinoid system in nature. Thus, the endogenous cannabinoid system, represented by cannabinoid receptors, endocannabinoids, and enzymes for the biosynthesis and degradation of these ligands, is conserved throughout evolution (Salzet et al., 2000). Endocannabinoids are present in peripheral and in brain tissues and have recently been proved to be present in breast milk (Di Marzo et al., 1998b). The expression of functional CB1Rs in the preimplantation embryo and synthesis of AEA in the pregnant uterus of mice suggest that cannabinoid ligand–receptor signaling is operative in the regulation of preimplantation embryo development and implantation (Paria and S.K., 2000). As the information on noladin ether, virodhamine, and NADA is limited, the detailed discussion is limited to AEA and 2‐AG.

2.2.1 Anandamide AEA was the first endocannabinoid substance to be isolated and structurally characterized (Devane et al., 1992). It was named anandamide from the Sanskrit ananda, ‘‘internal bliss,’’ and in reference to its chemical structure (the amide of arachidonic acid and ethanolamine) (> Figure 14-1). Later, ethanolamides of other fatty acids such as dihomo‐g‐linolenoylethanolamide and 7,10,13,16‐docosatetraenoylethanolamide, were included in the ‘‘anandamide family’’ (Hanus et al., 1993). AEA has been identified and quantitated throughout the human brain and periphery (Felder et al., 1996). Signal transduction and ligand binding studies have suggested that it can act at both the CB1 (Ki ¼ 61 nM) and the CB2 (Ki ¼ 1930 nM) receptors, although it may be more efficacious at CB1 (Felder et al., 1995). Some structural aspects required for AEA to bind to CB1Rs (Mechoulam et al., 1998) are (a) the number of double bonds on the fatty acid moiety has to be at least three or four; two double bonds only lead to in activation; (b) with or without hydroxylation on one of the alkyl groups, leads to loss of activity; (c) hydroxylation of the N‐monoalkyl group at the o‐carbon atom preserves activity; (d) the methyl ether and the phosphate are less active than the parent alcohol and the carboxylic acid derivatives are inactive; (e) the OH group in AEA can be replaced by fluorine with about tenfold increase in specific binding to CB1; and (f) conjugation of the double bonds leads to reduced activity. Some elementary structure–activity relationships such as the minimal and maximal length of the fatty acid residue; the effect of trans in place of cis double bonds; the conversion of the fatty acid chain into a prostaglandin‐type chain remains to be fully characterized. AEA also resembles D9‐THC (the main psychotropic constituent of marijuana) in structural aspects. The affinity of AEA for CB1Rs increases when its noncarboxylic hydrocarbon tail is lengthened and branched. As for the alkyl side chain of D9‐THC, extension of the noncarboxylic acid tail of AEA by two carbon atoms together with the introduction of two methyl substituents, produces an increase in both CB1‐ binding affinity and in vivo potency for CB1R‐mediated effects (Ryan et al., 1997; Seltzman et al., 1997). The AEA behaves as an affinity‐driven CB1R agonist. Thus its efficacy at CB1Rs, although higher than that of D9‐THC, is often found to be lower than that of other cannabinoid agonists [(þ)‐WIN55212‐2 or CP55940]. These similarities between D9‐THC and AEA are in line with the observations made in molecular‐ modeling studies. It was suggested that it is possible to superimpose AEA on the D9‐THC molecule such that the oxygen of the arachidonyl carboxyamide lies over the pyran oxygen, the hydroxyl group of the arachidonylethanol over the phenolic hydroxyl group, the five terminal arachidonyl carbons over the hydrophobic pentyl side chain, and the arachidonyl polyolefin loop over the tricyclic ring system (Thomas et al., 1996).

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2.2.1.1 Localization and Distribution of N‐Arachidonylethanolamine in Brain AEA was isolated and quantified by liquid chromatography and mass spectrometry in rat and postmortem human tissues. AEA was shown to be widely distributed in the brain and peripheral tissues (Felder et al., 1993; 1996). AEA levels vary by a factor of four to six within the different regions of the rat brain, with the highest levels in striatum and brain stem and the lowest levels in cerebellum and cortex (Bisogno et al., 1999; Yang et al., 1999). AEA was found in regions of both rat and human brain that contain a high density of CB1Rs (e.g., hippocampus, cerebellum, and striatum) and in a region that is sparse in CB1Rs, the thalamus (Felder et al., 1996). In rat, the concentration of AEA in the thalamus was approximately twice the concentration measured in the cerebellum. It is clear from these data that for AEA, the relative regional abundance in the brain does not correlate with the distribution of CB1Rs. The AEA levels in brain are equivalent to those of other neurotransmitters such as dopamine and serotonin, but at least tenfold lower than those reported for GABA and glutamate. AEA was also found in human and rat spleen, which expresses high levels of the CB2 receptors, suggesting that it is an agonist at both the CB1 and the CB2 receptor (Felder et al., 1996). Small amounts of AEA were also found in human heart and thalamus and also in rat skin, whereas only trace quantities were detected in human serum, plasma, and cerebrospinal fluid (Felder et al., 1996). 2.2.1.2 Biosynthesis and Metabolism of N‐Arachidonylethanolamine Soon after the discovery of AEA, enzymatic activities responsible for its biosynthesis and degradation were described. Unlike classical neurotransmitters and neuropeptides, AEA is not stored in intracellular compartments, but is produced on demand by receptor‐stimulated cleavage of lipid precursors (Di Marzo et al., 1994; Cadas et al., 1997; Mechoulam et al., 1998; Basavarajappa and Hungund, 1999a; Basavarajappa et al., 2000, 2003) and released from neurons immediately afterwards (Di Marzo et al., 1994; Mechoulam et al., 1998; Giuffrida et al., 1999; Basavarajappa and Hungund, 1999a; Basavarajappa et al., 2000, 2003). The AEA precursor is an N‐arachidonylphosphatidylethanolamine (N‐ArPE), which is believed to originate from the transfer of arachidonic acid from the sn‐1 position of 1,2‐sn‐di‐arachidonylphosphatidylcholine to phosphatidylethanolamine, catalyzed by a calcium‐dependent transacylase (CDTA) (> Figure 14-5). NArPE is then cleaved by a N‐acylphosphatidylethanolamine (NAPE)‐specific phospholiapse D (PLD) (Natarajan et al., 1981; Schmid et al., 1983; Di Marzo et al., 1994), which releases AEA and phosphatidic acid. It is not clear whether the N‐acyltransferase or the NAPE‐specific PLD controls the rate‐limiting step of AEA synthesis (> Figure 14-5) (Di Marzo, 1998; Hansen et al., 2000; Sugiura et al., 2002). However, the CDTA enzyme has since been characterized biochemically in both brain and testis microsomal preparations (Di Marzo et al., 1994; Sugiura et al., 1996a, b; Cadas et al., 1997). The CDTA has only recently been purified, cloned, and characterized (Okamoto et al., 2004). Another postulated biosynthetic pathway for AEA involves the energy‐independent condensation of ethanolamine and free arachidonic acid (Deutsch and Chin, 1993; Ueda et al., 1995). The biochemical characters and inhibitor kinetics observed for the enzyme catalyzing this condensation reaction were essentially identical to those described for the enzyme catalyzing fatty acid amide (FAA) hydrolysis [fatty acid amide hydrolase (FAAH); see discussion later] (Deutsch and Chin, 1993; Ueda et al., 1995; Cravatt et al., 1996). Several lines of evidence derived from both physical and biological data argue that the production of AEA in vivo does not occur through the energy‐independent condensation of free arachidonic acid and ethanolamine (Di Marzo et al., 1994; Kurahashi et al., 1997; Katayama et al., 1999). As a putative neuromodulator, AEA that is released into the synaptic cleft is expected to be rapidly inactivated. In general, two possibilities are known to remove a transmitter from the synaptic cleft to ensure rapid signal inactivation either reuptake or enzymatic degradation. An enzyme that catalyzes hydrolysis of AEA, forming arachidonic acid and ethanolamine, was detected in rat brain tissue (Deutsch and Chin, 1993) soon after the discovery of AEA and is variously known as ‘‘anandamide amidase,’’ ‘‘anandamide hydrolase,’’ ‘‘anandamide amidohydrolase,’’ or ‘‘FAAH’’ (see > Section 2.3 for detailed discussion). The existence of a membrane transporter that mediates the uptake of AEA has also been investigated (see > Section 2.4 for detailed discussion). Thus, the ability of brain tissue to enzymatically synthesize and metabolize AEA and also the existence of carrier‐mediated transport mechanisms essential for termination of the signaling effects of AEA and of specific receptors for the AEA suggest the presence of AEA‐containing (anandaergic) neurons. Therefore, AEA may represent a member of a new family of fatty‐acid‐derived neuromodulators (Ameri, 1999).

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. Figure 14-5 The mechanism of 2‐arachidonylglycerol formations. Intracellular Ca2þ initiates 2‐AG synthesis by inducing the formation of diacylglycerol (DAG) in the membrane either by stimulating the phosphatidyl‐inositol‐ phospholipase C (PI‐PLC) pathway or the formation of phosphatidic acid (PA), and the subsequent activation of phosphatidic acid hydrolase. 2‐AG is the product of DAG‐lipase acting on DAG. The second pathway involves hydrolysis of PI by phospholipase A1 (PLA1) and hydrolysis of the resultant lyso‐PI by a specific lyso‐PLC. Once synthesized, 2‐AG can be transported to the outside of the cell through a process that is not yet been characterized. 2‐AG is taken up from the extracellular space by a transport process that can be blocked by AM404, and is inactivated by fatty acid amide hydrolase (FAAH), although evidence that monoglyceride lipase may play a major role in 2‐AG inactivation has recently been reported (Dinh et al., 2002). PLA1, phospholipase A1; AMT, anandamide membrane transporter. For details, see > Section 2.2.2. [Reproduced with permission from Balapal Basavarajappa (2006)]

2.2.2 2‐Arachidonylglycerol Soon after the AEA discovery and cloning and the molecular characterization of CB1Rs, the hypothesis of the existence of an endogenous cannabinoid system was extended forward for the first time. A second phospholipid‐derived messenger, 2‐AG, has been identified and reported to display properties of an endogenous cannabinoid agonist (Stella et al., 1997). 2‐AG has been characterized as a unique molecular species of the monoacylglycerol isolated from canine gut (Mechoulam et al., 1995) and rat brain (Sugiura et al., 2000) as an endogenous cannabinoid receptor ligand. According to its chemical structure, this new putative endocannabinoid is an arachidonyl ester rather than an amide (> Figure 14-1); it was found to bind to both CB1 (Ki ¼ 2.4 mM) and CB2 receptors. The CB1R‐binding activity of 2‐AG was 24 times less potent than that of AEA. 2‐AG caused the typical effects of D9‐THC, such as antinociception, immobility, immunomodulation, and inhibition of electrically evoked contractions of the mouse vas deferens (Mechoulam et al., 1995; Sugiura et al., 1995, 1996c). However, further studies are necessary to determine the relative importance of 2‐AG in the human body and brain. Information regarding tissue levels of 2‐AG is still limited. Brain tissue concentrations of 2‐AG are approximately 200‐fold higher than those of AEA (Bisogno et al., 1999). The distribution of the two endocannabinoids in the different brain regions is similar. The highest concentrations were found in the

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brain stem, medulla, limbic forebrain, striatum, and hippocampus and the lowest in the cortex, diencephalons, mesencephalon, hypothalamus, and cerebellum (Sugiura et al., 2002). However, no correlation was found between 2‐AG concentrations and CB1R distribution. 2‐AG was also detected in the peripheral nervous system, such as in the sciatic nerve, lumbar spinal cord, and lumbar dorsal root ganglion. 2‐AG was also detected in the rat retina and bovine retina (Sugiura et al., 2002). These new research findings undoubtedly have advanced cannabis research and have allowed us to assume that the endocannabinoid system consists of a previously unrecognized elaborate network of endocannabinoid neurotransmitters complete with their accompanying biosynthetic, uptake, and degradation pathways. 2.2.2.1 Biosynthesis and Metabolism of 2‐Arachidonylglycerol Stimulus‐induced generation of 2‐AG was first described in ionomycin‐stimulated N18TG2 cells (Bisogno et al., 1997b), in electrically stimulated rat hippocampal slices, and in ionomycin‐stimulated neurons (Stella et al., 1997). Rapid generation of 2‐AG was also observed in rat brain homogenate during incubation with Ca2þ (Kondo et al., 1998) and in thrombin‐ or A23187‐stimulated human umbilical vein endothelial cells (Sugiura et al., 1998). Intraperitoneal injection of picrotoxin stimulated 2‐AG levels in rat brain (Sugiura et al., 2000). Generation of 2‐AG was also observed by chronic alcohol in cerebellar granular (CG) neurons in culture (Basavarajappa et al., 2000). Substantial amounts of this stimulus‐induced 2‐AG were released from the cells (Bisogno et al., 1997a; Sugiura et al., 1998; Basavarajappa et al., 2000). 2‐AG biosynthesis occurs by two possible routes in neurons, which are illustrated in > Figure 14-6. PLC‐mediated hydrolysis of membrane phospholipids may produce diacylglycerol (DAG), which may be subsequently converted to 2‐AG by diacylglycerol lipase (DGL) activity (Prescott and Majerus, 1983; Sugiura et al., 1995). Alternatively, phospholipase A1 (PLA1) may generate a lysophospholipid, which may be hydrolyzed to 2‐AG by lyso‐ PLC activity (Sugiura et al., 1995). Various structurally distinct inhibitors of PLC and DGL activities prevent 2‐AG formation in cultures of cortical neurons, suggesting that the PLC/DGL pathway may play a primary role in this process (Stella et al., 1997). The molecular character of the enzymes involved remains to

. Figure 14-6 Scheme illustrating the likely mechanism of anandamide uptake and degradation. Anandamide and 2‐arachidonylglycerol can be internalized by neurons through a yet unidentified transport mechanism, ‘‘the endocannabinoid transporter.’’ Once inside neurons, they can be hydrolyzed by hydrolytic enzyme and fatty acid amide hydrolase (FAAH). AMT, anandamide membrane transporter. For details, see > Sections 2.3 and > 2.4. [Reproduced with permission from Balapal Basavarajappa (2006)]

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be identified, although purification of rat brain DGL has been reported (Farooqui et al., 1986). Molecular characterization of these potential pathways remains to be accomplished. 2‐AG is inactivated by reuptake via a membrane transport molecule, the ‘‘AEA membrane transporter’’ (AMT) (see > Section 2.4 for details) (Beltramo et al., 1997; Hillard et al., 1997; Maccarrone et al., 1998; Beltramo and Piomelli, 2000; Hillard and Jarrahian, 2000; Giuffrida et al., 2001; Basavarajappa et al., 2003), and subsequent intracellular enzymatic degradation (Di Marzo et al., 1994; Day et al., 2001; Deutsch et al., 2001) by monoacylglycerol lipase, like other monoacylglycerols (Konrad et al., 1994). Similarly, 2‐AG is metabolized by monoacylglycerol lipase from porcine brain cytosol and particulate fractions (Goparaju et al., 1998). FAAH (for details see > Section 2.3) is also shown to metabolize 2‐AG (Goparaju et al., 1998; Di Marzo et al., 1998a). 2‐AG may be degraded by FAAH in addition to monoacylglycerol lipase under some circumstances. AEA and 2‐AG are inactivated by reuptake via a membrane transport molecule, the AMT (Beltramo et al., 1997; Hillard et al., 1997; Maccarrone et al., 1998; Beltramo and Piomelli, 2000; Hillard and Jarrahian, 2000; Giuffrida et al., 2001; Basavarajappa et al., 2003), and subsequent intracellular enzymatic degradation (Di Marzo et al., 1994; Day et al., 2001; Deutsch et al., 2001) by FAAH‐mediated hydrolysis (> Figure 14-7) (Cravatt et al., 1996; Beltramo and Piomelli, 2000; Ueda et al., 2000; Bisogno et al., 2001; Deutsch et al., 2001; Fowler et al., 2001). The distribution of FAAH and AMT in the brain is similar to that of the CB1 receptor. High concentrations are found in hippocampus, cerebellum, and cerebral cortex . Figure 14-7 Scheme illustrating the CB1 receptor signaling systems. CB1 receptors are 7‐transmembrane domain and G‐protein‐coupled located in the cell membrane. The Ca2þ channels inhibited by CB1 receptors include N‐, P/Q‐, and L‐type channels. Actions of GIRK and PI3K are thought to be mediated through bg subunits of the G‐protein, and those on Ca2þ channels and adenylyl cyclase (AC) by a subunits. Inhibition of AC and subsequent decrease in cAMP decreases activation of cAMP‐dependent protein kinase (PKA), which leads to decreased phosphorylation of the Kþ channels shown. Stimulatory effects are shown by (þ) sign and inhibitory effects by (
) sign. For details, see > section 2.1.7. [Reproduced with permission from Balapal Basavarajappa (2006)]

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(Egertova et al., 1998; Tsou et al., 1998b; Ueda et al., 2000; Giuffrida et al., 2001; Egertova et al., 2003). These observations conclusively support the hypothesis that the endocannabinoids are true neurotransmitters (Self, 1999).

2.3

Fatty Acid Amide Hydrolase

The known endocannabinoids, derived from membrane phospholipids that contain arachidonate, are metabolized by FAAH (Deutsch and Chin, 1993) (> Figure 14-7), which is a membrane‐associated serine hydrolase enriched in brain and liver. FAAH is a membrane‐associated enzyme that was first isolated from rat liver because of its ability to catalyze hydrolysis of the sleep‐inducing lipid oleamide (Cravatt et al., 1995, 1996). Analysis of the substrate selectivity of FAAH revealed, however, that it also hydrolyzes AEA, a putative endogenous ligand for the CB1‐type cannabinoid receptor in the brain (Cravatt et al., 1996). 2‐AG, a second putative endocannabinoid, also appears to be reactive with FAAH in vitro (Goparaju et al., 1998), and thus these data suggest that FAAH is generally required for inactivation of endocannabinoids in the brain. The actual enzymes involved in FAA metabolism remained unknown until the late 1990s, when a rat oleamide hydrolase activity was affinity purified and its cDNA cloned (Cravatt et al., 1996). This enzyme when recombinantly expressed, was shown to hydrolyze oleamide, AEA, and several other endogenous FAAs. Understanding of the three‐dimensional structure of enzymes can greatly assist inhibitor design (Rowland, 2002). Recently, the first X‐ray crystal structure of FAAH was reported (Bracey et al., 2002). This 2.8 A˚ structure of FAAH in complex with the irreversible inhibitor methoxy arachidonyl fluorophosphonate revealed several unusual features of the enzyme. First, the core catalytic machinery of FAAH is composed of a serine‐serine‐lysine catalytic triad (S241‐S217‐K142), in contrast to the serine‐histidine‐ aspartate triad typical of most serine hydrolases. These results are consistent with previous enzymological studies indicating that S241 and K142 play key roles in catalysis as the nucleophile and an acid/base, respectively (Patricelli et al., 1999). The structure of FAAH also revealed that this enzyme possesses a remarkable collection of channels that appear to grant it simultaneous access to both the membrane and cytoplasmic compartments of the cell, possibly to facilitate substrate binding, product release, and catalytic turnover. These unusual mechanistic and structural features of FAAH should inspire new strategies for the design of inhibitors that might display high selectivity for this enzyme relative to the hundreds of serine hydrolases present in the human proteome. Despite biochemical and cell biological studies supporting a role for AEA as an endogenous CB1 agonist, the behavioral effects elicited by the AEA are very weak and transient compared with those produced by D9‐THC (Adams et al., 1998). The limited pharmacological activity of AEA may be due to its rapid catabolism in vivo, as this lipid is hydrolyzed to arachidonic acid within minutes of exogenous administration (Willoughby et al., 1997). Nonetheless, the relative contribution made by FAAH to the hydrolysis of anandamide in vivo remained largely unclear until it was cloned and a mouse model was generated in which this enzyme was genetically deleted (FAAH knockout) (Cravatt et al., 2001). FAAH‐KO mice were shown to be born at the expected Mendelian frequency and were viable, fertile, and normal in their general cage behavior. Tissues from FAAH‐KO mice were found to display a 50–100‐fold reduction in hydrolysis rates for AEA and related FAAs. In contrast to FAAH wild‐type mice, in which administered AEA failed to produce significant behavioral effects, FAAH‐KO mice exhibited robust responses to AEA, becoming hypomotile, analgesic, cataleptic, and hypothermic. Further, all of the behavioral effects of AEA in FAAH‐KO mice were blocked by pretreatment with the SR141716A, indicating that AEA acted as a potent and selective CB1 agonist in these animals. Consistent with this notion, the brain homogenates from FAAH‐KO mice showed approximately 15‐fold higher apparent binding affinity of AEA for the CB1R (Lichtman et al., 2002). These FAAH‐KO mice have been shown to have elevated (10–15‐fold) levels of AEA and other NAE in several brain regions, including hippocampus, cortex, and cerebellum (Cravatt et al., 2001; Clement et al., 2003). Interestingly, these increased levels of AEA and NAEs in the brain correlated with a CB1‐dependent reduction in pain sensation in FAAH‐KO mice (Cravatt et al., 2001). These observations taken together suggest that FAAH is a key enzyme involved in catabolism of AEA and other NAEs in vivo and suggest that

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pain pathways are under the influence of a FAAH‐regulated endocannabinoid tone. However, it should be noted here that FAAH‐KO mice exhibited normal motility, body weight, and body temperature, suggesting that several other neurobehaviors affected by exogenously applied CB1 agonists were not under tonic control of AEA in these animals.

2.3.1 Localization and Distribution of Fatty Acid Amide Hydrolase in the Brain The majority of FAAH in the mouse brain is associated with neurons; this is particularly interesting and probably most directly relevant to endocannabinoid signaling. FAAH appears not to be confined to neuronal somata but also extends into surrounding areas of neural tissue. This is most clearly seen in the hippocampal formation where although the highest concentration of FAAH is clearly located in the somata of hippocampal pyramidal cells and granule cells of the dentate gyrus, there is also widespread presence of FAAH in adjacent layers that contain their axons and dendrites (Egertova et al., 2003). The most striking and intensely stained FAAH‐immunoreactivity neurons were found in the mesencephalic trigeminal nucleus (Me5) (Egertova et al., 2003). In addition to neuronal FAAH expression, FAAH is also expressed by oligodendrocytes and ventricular ependymal cells of the mouse brain (Egertova et al., 2003). The functional significance of FAAH expression in these non‐neuronal cells is not yet known. FAAH‐expressing neurons were also found in brain regions (thalamus, midbrain, and hind brain) known to express few or no CB1Rs. However, there are clearly some regions of the brain (e.g., thalamic nuclei, mesencephalic trigeminal nuclei, cerebellar nuclei) where FAAH‐expressing neurons are widespread but lack associated CB1 expression. In regions of the brain such as these, the role of FAAH is likely to be unrelated to CB1‐dependent endocannabinoid signaling mechanisms (Egertova et al., 2003). More typically, however, FAAH and CB1 are anatomically associated, with FAAH‐immunoreactive neuronal somata surrounded by CB1‐immunoreactive fibers, in the rat cerebellar cortex, hippocampus, and neocortex (Egertova et al., 1998). FAAH is likely to influence retrograde synaptic signaling (for detailed discussion see > Section 2.5) molecules, particularly in the cerebellar cortex, where endocannabinoids mediate retrograde suppression of both excitatory and inhibitory synapses onto Purkinje cells (Maejima et al., 2001; Kreitzer and Regehr, 2001a, b). Here FAAH is located in Purkinje cell somata, whilst CB1 is located on the presynaptic terminals of granule cells (glutamatergic parallel fibers), basket cells (GABAergic), and probably also climbing fibers (glutamatergic). Depolarization of Purkinje cells causes transient endocannabinoid‐mediated inhibition of excitatory inputs from parallel fibers and climbing fibers, with maximal inhibition occurring after approximately 5 s, followed by a gradual restoration of basal excitatory input within 50 s (Kreitzer and Regehr, 2001a). Because FAAH is located in the somato‐dendritic compartment of Purkinje cells, the duration of the inhibitory action of Purkinje cell–derived endocannabinoids is likely to be largely influenced by uptake and FAAH‐mediated inactivation of endocannabinoids in Purkinje cells (Egertova et al., 2003). FAAH is also present in fibers of the olfactory nerves that project into the olfactory bulb glomeruli where they form synapses with the glomerular dendrites of mitral cells. This is of particular interest because FAAH is targeted to the axonal compartment of neurons because in all other regions of the mouse and rat brain it is targeted to the somato‐dendritic compartment of neurons. It also provides a unique example of a synapse where FAAH is located both presynaptically (olfactory‐receptor neuron terminals) and postsynaptically (mitral cells) (Egertova et al., 2003). The functional significance of this complex pattern of expression is not yet clear. The distribution of CB1 in the brain, although strikingly complementary with FAAH in many regions of the forebrain and in the cerebellar cortex, is not always associated with FAAH‐expressing neurons. For example, there is a population of striatal GABAergic medium‐spiny projection neurons that target the CB1R to their axonal terminals in globus pallidus, entopeduncular nucleus (EP), and SNR. The density of CB1Rs in these output nuclei is particularly high, whilst FAAH‐expressing neurons are sparse or absent (Egertova et al., 2003).

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A naturally occurring single nucleotide polymorphism in the human FAAH gene, 385A, was strongly associated with street drug use and problem drug/alcohol use. This association seems especially relevant to illegal drug use, because neither alcohol nor nicotine abuse alone showed any significant relationship to the 385C‐A missense mutation. The association of a naturally occurring human FAAH mutation with problem drug use provides further support that the endogenous cannabinoid system plays an important role in neural circuits that underlie drug abuse and dependence (Sipe et al., 2002) including alcohol (Basavarajappa and Hungund, 2002; Basavarajappa et al., 2003).

2.4

Endocannabinoids Uptake

AEA appears to be produced from phospholipids in a stimulus‐dependent manner by the consecutive action of two enzymes, a calcium‐dependent transacylase and a phospholipase D (discussed in > Section 2.2.1.2), and inactivated by a two‐step process involving the transport of this lipid into cells (Beltramo et al., 1997) (> Figure 14-7) followed by intracellular hydrolysis by the integral membrane enzyme FAAH (discussed in > Section 2.3). However, with respect to the inactivation of AEA, only FAAH has been molecularly characterized and structurally studied. Indeed, the actual nature and mechanism of AEA uptake or, more generally, the movement of this FAA not only through biological membranes but also through aqueous media (extracellular and cytoplasmic) remains an enigmatic and controversial subject as to the nature of this uptake process. Some authors have suggested that the uptake is due to a facilitated diffusion process (Hillard et al., 1997; Basavarajappa et al., 2003), whereas others have described it as a passive process driven by the anandamide metabolizing enzyme FAAH (Day et al., 2001; Deutsch et al., 2001; —for a current discussion of these two points of view, see Glaser et al., 2003; Hillard and Jarrahian, 2003). AEA uptake by cells occurs via diffusion through the cell membrane, facilitated by a saturable, temperature‐dependent, and selective transport system (Di Marzo et al., 1994). Such a transporter, the AMT, has been identified in most of the cells analyzed so far (Hillard and Jarrahian, 2000) (> Figure 14-7), and inhibitors capable of enhancing AEA actions in vitro and in vivo have been developed (Beltramo et al., 1997; Di Marzo et al., 1998c). In addition, structure–activity relationship studies have been carried out on the AMT with a large variety of AEA analogs (Melck et al., 1999; Piomelli et al., 1999; Hillard and Jarrahian, 2000; Jarrahian et al., 2000; Palmer et al., 2000; Di Marzo et al., 2000a). It was established that at least one to four cis double bonds in the fatty acyl chain are necessary in order to simply bind to the AMT or be transported into cells (Melck et al., 1999). On the other hand, it was also observed that the ethanolamine ‘‘head’’ of AEA could be substituted for more hindering groups, better if aromatic, and still yields compounds capable of binding to the AMT (Melck et al., 1999; Jarrahian et al., 2000). Finally, it was reported that the AMT is present in endothelial cells (Maccarrone et al., 2001), is activated by nitric oxide (NO) (Maccarrone et al., 1998, 1999, 2000a, b), and is inhibited by plant cannabinoids, THC, and cannabidiol at micromolar concentrations (Rakhshan et al., 2000). The role of intracellular catabolism of AEA in driving, in part, the AMT was also pointed out (Rakhshan et al., 2000; Deutsch et al., 2001). The uptake of 2‐AG by cells was first observed in rat basophilic RBL‐2H3 and mouse neuroblastoma N18TG2 cells, and was shown to be inhibited by unsaturated monoacylglycerols such as 2‐oleoyl‐, 2‐linoleoyl‐, and 2‐linolenoylglycerols (Ben‐Shabat et al., 1998). At a closer look, it was observed that although taken up in a temperature‐dependent manner unmetabolized 2‐AG, unlike AEA, would not greatly accumulate in cells and, in any case, not in a fashion sensitive to low temperature (Di Marzo et al., 1996; 1998a; 1999; Maccarrone et al., 2000a). In studies carried out in RBL‐2H3 and J774 cells (Di Marzo et al., 1996, 1998a; Rakhshan et al., 2000) it was also observed that 2‐AG does not inhibit efficiently the uptake by cells of [14C]AEA when the two substances are present at the same concentration, thus suggesting that the AMT would not recognize 2‐AG as a substrate. Very recently, a study carried out in human astrocytoma cells (Beltramo and Piomelli, 2000) confirmed that 2‐AG is taken up by cells, and showed for the first time that this process can be inhibited by the previously developed AMT inhibitor, AM404 (Beltramo et al., 1997).

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Roles of Endocannabinoids in Central Nervous System

The discovery of endocannabinoids such as anandamide and 2‐AG and the widespread localization of CB1Rs in the brain suggest that cannabinoid system represents a previously unrecognized ubiquitous network in the nervous system, whose biology and role is unfolding. There is now overwhelming evidence that the anandamide and 2‐AG interact with the CB1Rs and share some of the biological properties of other cannabinoids with significant differences. These significant differential effects involve other non‐CB1R and/ or unknown CB3 receptors as described. Despite the decades of extensive investigations and recent developments in cannabinoid research, the identification of specific mechanisms for the actions of endocannabinoids has been slow to emerge. In the year 2001, we witnessed the functional roles of endocannabinoids at the synaptic and network levels. Therefore, an attempt has been made to provide a comprehensive account of the many physiological roles of endocannabinoids in the CNS.

2.5.1 Regulation of g‐Aminobutyric Acid Transmission It was reported that brief activation of CA1 pyramidal cells in the hippocampus (Pitler and Alger, 1990, 1992, 1994; Alger et al., 1996; Ohno‐Shosaku et al., 1998; Alger, 2002) or Purkinje cells in the cerebellum (Llano et al., 1991; Vincent et al., 1992; Vincent and Marty, 1993, 1996) (> Figure 14-8) caused a reduction . Figure 14-8 A hypothetical model to illustrate the role of AEA and CB1 receptors on excitatory and inhibitory neurotransmissions. Depolarization of postsynaptic neuron causes the generation and the release of endocannabinoids such as anandamide (AEA). The released endocannabinoids then activate the CB1 receptors (CB1R) at presynaptic terminals and suppress the release of glutamate (left) by inhibiting Ca2þ channels (Shen and Thayer, 1998; Hoffman and Lupica, 2000; Huang et al., 2001). Depolarization of postsynaptic neuron causes activation of CB1 receptors at GABAergic neurons and causes the inhibition of GABA release. For details, see > section 2.5. [Reproduced with permission from Balapal Basavarajappa (2006)]

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in the amplitude of GABAergic inhibitory postsynaptic currents (IPSCs). This phenomenon, called depolarization‐induced suppression of inhibition (DSI), is initiated postsynaptically by the voltage‐dependent influx of Ca2þ into the soma and dendrites of the neuron, but is expressed presynaptically through inhibition of transmitter release from axon terminals of GABA interneurons. This suggests that a chemical messenger generated during depolarization of the pyramidal neurons must travel backwards across the synapse to induce DSI. DSI has been observed in both excitatory and inhibitory neurons in hippocampal cell culture (Ohno‐Shosaku et al., 2000), in CA3 pyramidal cells (Morishita and Alger, 2000), in dentate gyrus granule cells (Alger, 2002), and in neocortical pyramidal cells (Zilberter, 2000). The retrograde messenger in DSI remained unknown until recent investigation reported by Wilson and Colleagues (Wilson and Nicoll, 2001, 2002; Ohno‐Shosaku et al., 2001; Wilson et al., 2001) that it was likely to be an endocannabinoid in hippocampal cells. Shortly thereafter, cerebellar DSI was also reported to be mediated by an endocannabinoid (Kreitzer and Regehr, 2001a; Diana et al., 2002; Yoshida et al., 2002). It was reported that CB1R agonists selectively reduced IPSCs in the hippocampus (Katona et al., 1999; Hajos et al., 2000; Hoffman and Lupica, 2000) and cerebellum (Takahashi and Linden, 2000). There is strong evidence that this retrograde‐signaling process involves an endocannabinoid. (a) The CB1R antagonists selectively blocked DSI, whereas agonists occluded it (Ohno‐Shosaku et al., 2001; Wilson and Nicoll 2001). (b) The DSI was absent in CB1R knockout mice (Wilson et al., 2001; Yoshida et al., 2002). (c) The GABA interneurons that are implicated in DSI express high levels of CB1Rs, which are localized to their axon terminals (Katona et al., 1999). (d) Neuronal activity and Ca2þ entry stimulate the synthesis of 2‐AG in hippocampal neurons and AEA and 2‐AG in other neuronal cells (Basavarajappa and Hungund, 1999a; Basavarajappa et al., 2000, 2003). It remains to be investigated whether endocannabinoid‐mediated DSI reported in other brain regions such as the ventromedial medulla (Vaughan et al., 1999), amygdale (Katona et al., 1999), substantia nigra (Wallmichrath and Szabo, 2002), and striatum (Szabo et al., 1998), in which exogenously applied CB1R agonists are known to suppress IPSCs.

2.5.2 Regulation of Glutamate Transmission Metabotropic glutamate receptors (mGluRs) are a family of GPCRs distributed throughout the CNS and modulate multiple CNS functions, including neuronal excitability (Conn and Pin, 1997; Anwyl, 1999) and neurotransmitter release (Cartmell and Schoepp, 2000). Recent data from several investigators have begun to uncover an entirely novel‐signaling mechanism for mGluRs, namely, the production and subsequent release of the endocannabinoids. Some of the effects (short‐ and long‐term forms of synaptic plasticity) previously attributed directly to mG1uR activation are in reality indirectly mediated by signaling through the endocannabinoid system (Doherty and Dingledine, 2003). Recent studies suggest that mG1uR/endocannabinoid signaling is a widespread feature of neuronal circuitry (Doherty and Dingledine, 2003), given the widespread expression of postsynaptic group I mG1uRs throughout the CNS, and a similar extensive expression of CB1Rs. Activation of group I mG1uRs can cause the release of endocannabinoids in the cerebellum (Maejima et al., 2001) and hippocampus (Varma et al., 2001). In the cerebellum, a selective mG1uR1 agonist, DHPG, reduced glutamate release and this effect was prevented by the mG1uR1 antagonist and was absent in mG1uR1 knockout mice. Despite the presynaptic locus of its effects, it was clear that the mG1uR action was exerted on the postsynaptic Purkinje cells, as injection of GTPgS into the Purkinje cell entirely abolished the effects of the bath‐applied DHPG. The effects of DHPG were prevented by the SR141716A or AM‐281, and occluded by WIN55212‐2. Synaptic depression in the cerebellum has been observed at glutamatergic synapses. A similar mechanism operates in the hippocampus. As in the cerebellum, activation of postsynaptic group I mG1uRs depresses neurotransmitter release through endocannabinoid‐mediated retrograde activation of presynaptic CB1Rs (> Figure 14-8). In contrast to the cerebellum, however, synaptic depression in the hippocampus has been observed at GABAergic rather than glutamatergic synapses. The neurons in the hippocampus and cerebellum use endocannabinoids to carry out a signaling process that is analogous in mechanism, but opposite in sign, to DSI, called depolarization‐induced suppression of excitation (DSE). Like DSI, DSE is induced by neuronal depolarization; it consists of transient depression in

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neurotransmitter release, and it requires a retrograde endocannabinoid messenger. However, unlike DSI, DSE targets glutamatergic rather than GABA axon terminals and results therefore in reduced excitatory input to the affected cell (Alger, 2002; Piomelli, 2003). DSE is mimicked and occluded by agonists and antagonists of CB1R (Maejima et al., 2001; Kreitzer and Regehr, 2001b) and it is absent in the CB1R knockout mouse (Ohno‐Shosaku et al., 2002). Interestingly, a novel type of cannabinoid receptor (CB3) was inferred as the WIN55212‐2, continued to suppress excitatory postsynaptic currents (EPSCs) in a CB1R knockout mouse (Hajos et al., 2001). In another study, hippocampal DSE, like DSI and the actions of WIN55212‐2 on both IPSCs and EPSCs, was reportedly absent in CB1R knockout mice (Ohno‐Shosaku et al., 2002), contrary to the findings of Hajos et al. (2001). This discrepancy was later related to differences in developmental stages, and further study will be required to resolve these issues (Alger, 2002). CB1R agonists suppress EPSCs in other areas of the brain, evidently through presynaptic actions. For instance, similar DSE was reported in the ventral tegamental area (VTA) as a Ca2þ‐dependent phenomenon, blocked by both AM281 and SR141716A, and occluded by WIN55212‐2 (Melis et al., 2004). Importantly, DSE was partially blocked by the D2 DA antagonist eticlopride and enhanced by the D2 DA agonist quinpirole without changing the presynaptic cannabinoid activity (Melis et al., 2004). These observations indicate that activation of D2 DA receptors in the VTA significantly enhances the depolarization‐induced release of endocannabinoids, which was responsible for the inhibition of glutamate transmission in the VTA (Melis et al., 2004). A synchronous release of mEPSCs in Sr2þ‐substituted extracellular solution could be reduced by endocannabinoids in the prefrontal cortex and striatum (Auclair et al., 2000; Gerdeman and Lovinger, 2001). It remains to be demonstrated whether or not DSE is present in the striatum (Gerdeman and Lovinger, 2001), substantia nigra (Szabo et al., 2000), periaqueductal gray (Vaughan et al., 2000), and spinal cord (Morisset and Urban, 2001).

2.5.3 Mechanism of Inhibition of Neurotransmitter Release A very important issue for understanding the functional aspects of endocannabinoids is how far they spread from their point of release. At present this is a somewhat controversial issue, with some studies showing clear evidence for spread (Wilson and Nicoll, 2001; Kreitzer and Regehr, 2001a), and others showing the opposite (Pitler and Alger, 1994; Maejima et al., 2001; Morishita and Alger, 2001). Another important issue remaining to be determined is whether or not DSI spreads to other cells under true physiological conditions. It is yet unclear how this endocannabinoid makes its way to the presynaptic nerve terminal, or is produced there by the action of another unknown signaling molecule. It will also be important to determine if retrograde actions of endocannabinoids (DSI‐ and DSE‐ inducing effects) affect release of other transmitters, including acetylcholine, norepinephrine, and glycine, whose release is influenced by exogenous cannabinoids (see Freund et al., 2003 for review). Numerous studies addressed the issue of how endocannabinoids affect transmitter release from presynaptic neurons (Alger, 2002). It is clearly established that the bg subunits of heterotrimeric G proteins directly interact with and inhibit the high voltage–activated Ca2þ channels that mediate transmitter release at most synapses (> Figure 14-8). CB1Rs are pertussis‐sensitive GPCRs, and their activation reduces N‐type Ca2þ current (Caulfield and Brown, 1992; Caulfield et al., 1992; Mackie and Hille, 1992; Pan et al., 1996; Twitchell et al., 1997). In some cells, it appears that the CB1R could reduce GABA release from at least some nerve terminals through a mechanism that is independent of N‐P/Q‐type Ca2þ channels (Takahashi and Linden, 2000; Vaughan et al., 2000; Gerdeman and Lovinger, 2001; Varma et al., 2001), perhaps a direct action on the transmitter release machinery.

2.5.4 Release of Endocannabinoids by Activation of Other Neurotransmitter Receptors Several studies showed that conventional neurotransmitters could induce the formation and/or release of endocannabinoids. Electrical stimulation of a glutamatergic fiber tract in hippocampal slices caused a significant, Ca2þ‐ and TTX‐sensitive increase in 2‐AG levels, without affecting AEA concentration (Stella

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et al., 1997). Activation of an NMDA receptor increased the levels of 2‐AG, but not of AEA, in rat cortical neurons (Stella and Piomelli, 2001). By contrast, in the same study, coactivation of nicotinic cholinergic receptors (nAChRs) and NMDA receptors was found to increase AEA levels. Activation of NMDA receptors increased the levels of 2‐AG in primary cultures of rat CG neurons (Basavarajappa et al., 2000). Activation of type 1 mG1u receptors caused the release of endocannabinoids in the hippocampus (Varma et al., 2001) and cerebellum (Maejima et al., 2001). It was shown that activation of muscarinic acetylcholine receptors (mAChRs) by carbachol enhances DSI and this effect was inhibited by a CB1R antagonist and was absent in the CB1R knockout mouse, indicating that they are mediated exclusively by endocannabinoids (Kim et al., 2002). Activation of D2‐like (D2–D4) receptors, but not D1‐like (D1 and D5) DA receptors caused marked increases in AEA levels in the dorsal striatum (Giuffrida et al., 1999). The D2 receptor antagonist haloperidol inhibited the alcohol‐induced formation of 2‐AG in CG neurons (Basavarajappa et al., 2000).

2.5.5 Endocannabinoids and Synaptic Plasticity AEA is demonstrated to be an effective inhibitor of new synapse formation, raising the interesting possibility that the endocannabinoid system regulates the number of functional synapses (Abel, 1970; Tart, 1970; Chaperon and Thiebot, 1999). It was observed that striatal long‐term depression (LTD) was absent in CB1R knockout mice, reduced or eliminated by the SR141716A, and occluded by the HU‐210. These data suggested that striatal LTD was mediated by an endocannabinoid (Gerdeman et al., 2002). As in the striatum, LTD in the NAc is prevented by CB1R antagonists, occluded by CB1R agonists, and absent in CB1R knockout mice (Robbe et al., 2002). Importantly, once NAc LTD was induced, the antagonist did not affect it, demonstrating that the LTD was not maintained by a continual release of endocannabinoids, but rather represented a persistent effect of transient CB1R activation. The endocannabinoid that mediated LTD was evidently released as a retrograde messenger, because LTD was prevented by chelating postsynaptic Ca2þ (20 mM BAPTA) in the recorded cell (Robbe et al., 2002). Although, the mechanism needs to be elucidated, these studies suggest that endocannabinoid actions are important for the induction, but not the expression, phase of LTD. CB1Rs are present on glutamatergic terminals in the prefrontal cortex (Herkenham et al., 1990), and activation of the CB1R by agonists suppresses glutamate EPSCs in layer V slices of rat cortex, evidently by acting at a presynaptic site (Auclair et al., 2000), and it remained to be determined whether CB1Rs mediate LTD in cortex as well. It is known that LTP and LTD of CA3–CA1 synaptic transmission are two in vitro models for learning and memory. It has been shown that CB1R activation inhibits both LTP and LTD induction in the hippocampus (Stella et al., 1997; Sullivan, 2000). The mechanism by which cannabinoids inhibit LTP and LTD needs to be worked out. This may provide a clue to the cellular and molecular mechanisms underlying some of the cannabinoid‐induced learning and memory impairments. Another important finding is the enhanced LTP (Bohme et al., 2000) found in CB1R knockout mice. These data could not be mimicked by application of CB1R antagonists to wild‐type mice (Marsicano et al., 2002), suggesting some other factor in the enhancement of LTP. Compensatory upregulation of other genes in the CB1R knockout mice or altered neurophysiological responses might be responsible. An alternative explanation might involve a loss of CB1R cholinergic or noradrenergic afferents to the hippocampus resulting in enhanced release of these transmitters, which may contribute to the facilitation of LTP (Stanton and Sarvey, 1985; Brocher et al., 1992; Katsuki et al., 1997; Kobayashi et al., 1997). These early investigations just begin to address the effects of endocannabinoids on the neurophysiology of the brain, and further studies need to be done before their roles are fully elucidated.

2.5.6 Role of Endocannabinoid System in Disease The involvement of the endocannabinoid system in several neurodegenerative diseases has been reviewed recently (Glass, 2001; Drysdale and Platt, 2003). Impaired levels of endocannabinoids, CB1Rs, and CB1R mRNA have been reported in Huntington’s disease (Denovan‐Wright and Robertson, 2000; Glass et al., 2000, 2004). Although the mechanism and significance of the loss of endocannabinoids and CB1R levels are

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not clear at present, these observations may indicate that the endocannabinoid system has a central role in the progression of neurodegeneration in Huntington’s disease. Increased cannabinoid tone in the globus pallidus has been reported to be responsible for the production of Parkinsonian symptomology (Maneuf et al., 1996). A recent study demonstrated increased 2‐AG in the globus pallidus of rat treated with resperpine, a rodent model of Parkinson’s disease (Di Marzo et al., 2000b). The deficiency in endocannabinoid transmission may contribute to levodopa‐induced dyskinesias; these complications may be alleviated by activation of CB1Rs (Ferrer et al., 2003). Several lines of evidence exist to suggest a role for endocannabinoid signaling in schizophrenia (Glass, 2001). CB1Rs of highest densities are found in regions of the human brain that have been implicated in schizophrenia, including the prefrontal cortex, basal ganglia, hippocampus, and anterior cingulate cortex (Glass, 2001). Increased binding of the [3H]CP55940 to CB1Rs in the dorsolateral prefrontal cortex of schizophrenia patients compared to controls was shown (Dean et al., 2001). In addition, Leweke et al. (1999) reported significant twofold elevations of AEA levels in the cerebrospinal fluid of patients with schizophrenia compared to age‐matched controls. Finally, a recent study also indicates that SR141716A reverses ketamine‐induced impairment in prepulse inhibition of the acoustic startle reflex, an animal model of the deficient sensorimotor gating observed in schizophrenia (Musty et al., 2000). AEA release in the dorsal striatum is stimulated by activation of D2 dopamine receptors (Ferrer et al., 2003). Blockade of CB1Rs by SR141716A improves amnesia induced by a b‐amyloid fragment in mice, suggesting that the endocannabinoid system may be involved in cognitive impairment in Alzheimer’s disease (Mazzola et al., 2003). It is interesting to note the transient inhibition of food intake and reduction in fat mass that is observed following treatment of mice and rats with the SR141716A (Ravinet Trillou et al., 2003). The CB1R knockout mice have been shown to have lower susceptibility to obesity following a high‐fat diet (Ravinet Trillou et al., 2004). These observations suggest that the CB1R may have a role in obesity. Psychopathological disorders, and depression in particular, are strongly linked to eating attitude in obese patients. The identification of cannabinoid CB1Rs in areas of the CNS that have been implicated in regulation of mood and food intake suggests that these receptors may mediate such a behavioral link. A study implicates enhanced cannabinoid receptor and its function in the postmortem prefrontal cortex of depressed suicide brain (Hungund et al., 2004). In recent years, experimental evidence has suggested that the endocannabinoid system is a major player among the neurotransmitter systems involved in the regulation of different alcohol‐related phenomena including tolerance, vulnerability, reinforcement, and consumption (for more details, see recent reviews) (Hungund and Basavarajappa, 2000a, b, 2004; Basavarajappa and Hungund, 2002, 2005; Hungund et al., 2002; Basavarajappa, 2006). It was demonstrated that AEA and 2‐AG synthesis was increased by chronic alcohol. Further, chronic alcohol treatment led to a significant increase in the brain levels of AEA and a significant reduction in N‐arachidonyl phosphatidyl ethanolamide (NArPE), an immediate precursor for AEA synthesis, compared to control brains (Hungund et al., 2002). Chronic alcohol exposure of rats caused a decrease in the content of both AEA and 2‐AG in the midbrain, whereas AEA content increased in the limbic forebrain, a key area for the reinforcing properties of habit‐forming drugs including alcohol (Gonzalez et al., 2002). These observations indicate the possible involvement of the endocannabinoids in alcohol‐induced neuroadaptive changes in brain and that activation of endocannabinoid‐mediated neurotransmission may be responsible for the activation of the reward system by alcohol. We found that chronic exposure to alcohol leads to increase in extracellular AEA by inhibiting the uptake of AEA. This effect was independent of the CB1R as CB1R knockout mice have normal uptake activity (Basavarajappa et al., 2003). After prolonged exposure to alcohol, cells become tolerant to this effect such that AEA uptake is no longer inhibited by acute alcohol (Basavarajappa et al., 2003). Chronic alcohol did not show any direct inhibition of FAAH activity in these neurons. These data suggest that alcohol‐induced inhibition of AEA uptake; may in part, be responsible for alcohol‐induced increase in extracellular AEA. Evidence demonstrating the downregulation of CB1R numbers and function in chronic alcohol‐ exposed mouse brain has recently been reported. The results of this study indicate that chronic alcohol exposure decreased the Bmax of CB1Rs (without any changes in the receptor affinities) (Basavarajappa et al., 1998). These observations are consistent with the recent data which indicate that forced consumption of high levels of alcohol significantly decreased CB1R gene expression in the caudate putamen (Cpu), ventromedial nucleus of the hypothalamus (VMN), CA1 and CA2 fields of the hippocampus (Ortiz

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et al., 2004a). Recent studies to examine if the chronic alcohol‐mediated downregulation of brain CB1Rs has any functional effect on CB1R‐activated G proteins revealed that the net CB1R agonist (CP55940)‐ stimulated [35S] GTPgS binding was reduced significantly in chronic alcohol‐exposed mice without any significant changes in the G‐protein affinity (Basavarajappa and Hungund, 1999b). These results strongly support the participation of the endocannabinoid‐signaling mechanism in mediating some of the pharmacological and behavioral effects of alcohol, and hence the CB1R may constitute an important target for therapeutic intervention in alcohol‐related behaviors. There is strong evidence that the dopaminergic system that projects from the VTA of the midbrain to the NAc, and to other forebrain sites including the dorsal striatum, is the major substrate of reward and reinforcement produced by most drugs of abuse including alcohol (Wise and Bozarth, 1987; Di Chiara and Imperato, 1988; Robbins and Everitt, 1996; Wise 1996). It is well established that cannabinoids activate dopaminergic neurons in the VTA (Wise and Bozarth, 1987; Di Chiara and Imperato, 1988; Robbins and Everitt, 1996; Wise 1996; Tanda et al., 1997; Gessa et al., 1998), resulting in the release of DA in the NAc (Szabo et al., 1999). Recent results provide unequivocal evidence that the acute alcohol‐induced DA release in NAc is in fact mediated by CB1Rs (Hungund et al., 2003). The acute alcohol‐induced increase in DA in NAc dialysates in C57BL/6 mice was completely inhibited by pretreatment with SR141716A or deletion of CB1Rs in mice (CB1R knockout) (Hungund et al., 2003). These observations indicate that activation of the limbic DA system is required for the reinforcing effects of alcohol and further suggest an interaction between the cannabinoidergic and dopaminergic systems in the reinforcing properties of alcohol abuse. Several studies have shown the inhibition of voluntary alcohol intake by SR141716A in rodents. SR141716A has been shown to decrease voluntary alcohol intake in alcohol‐preferring C57BL/6 mice (Arnone et al., 1997), in Sardinian alcohol‐preferring (sP) rats (Colombo et al., 1998), in alcohol self‐administering Long Evans rats (Freedland et al., 2001), and in alcohol‐preferring congenic B6.Cb4i5‐b/13C/Vad and B6. Cb4i5‐b14/Vad mouse strains (Hungund et al., 2002). The acute administration of SR141716A suppressed alcohol self‐administration in chronic alcohol‐exposed Wistar rats (Rodriguez de Fonseca et al., 1999). A significant increase in alcohol preference (free choice) was observed when Wistar rats were treated with acute dose of SR141716A during chronic alcohol treatment (Lallemand et al., 2001). The administration of SR141716A after chronic alcoholization significantly decreased the preference for alcohol. Alcohol withdrawal symptoms were also decreased by administration of SR141716A in these studies. Furthermore, acute administration of the CB1R agonist CP55940 increased the motivation to consume alcohol in Wistar rats, and this effect was completely prevented by pretreatment with the CB1R antagonist SR141716A (Gallate and McGregor, 1999; Gallate et al., 1999). An acute dose of SR141716A completely abolished the alcohol deprivation effect (i.e., the temporary increase in alcohol intake after a period of alcohol withdrawal) in sP rats (Serra et al., 2002). Acute administration of the CB1R agonists WIN55212‐2 and CP55940 significantly stimulated voluntary alcohol consumption in alcohol‐preferring sP rats and this was completely prevented by SR141716A (Colombo et al., 2002). The existing evidence suggests that activation of the CB1R by a CB1R agonist may involve the release of DA in the NAc and inactivation of the CB1R by SR141716A may inhibit the DA release. This in turn may regulate voluntary alcohol intake in these animals. Evidence to show the participation of the cannabinoidergic system in alcohol‐drinking behavior is derived from the observed differences in the CB1R function in two genetic strains of alcohol‐preferring C57BL/6 and alcohol‐avoiding DBA/2 mice. In this study, it was found that C57BL/6 mice have a significantly lower level of CB1R‐binding sites and higher affinity for [3H] CP55940 than DBA/2 mice (Hungund and Basavarajappa, 2000c). Interestingly, the significantly higher levels of CB1Rs found in DBA/ 2 mice are less coupled to G proteins as shown by GTPgS‐binding assay compared to C57BL/6 mouse strains (Basavarajappa and Hungund, 2001), suggesting the participation of these receptors in controlling voluntary alcohol consumption. Recent study shows the lower regional level of CB1R function and lower CB1R gene expression in the brain of Fawn Hooded (alcohol‐preferring) than in Wistar rats (alcohol‐ nonpreferring) (Ortiz et al., 2004b). These results further strengthen the hypothesis that the inactivation or downregulation of CB1Rs could block voluntary alcohol consumption. It is therefore quite possible that the animals with a genetic predisposition to high alcohol consumption have inherited a greater sensitivity of the endogenous cannabinoidergic system to alcohol. This hypothesis was further examined using genetically modified CB1R knockout mice. CB1R knockout mice exhibited dramatically reduced voluntary alcohol

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consumption (Hungund et al., 2003). There was also a remarkable gender difference, with female wild‐type mice consuming significantly more alcohol than male wild‐type mice, whereas this gender difference was nonexistent in male and female CB1 knockout mice (Hungund et al., 2003). These results strongly support our hypothesis that the CB1R system plays an important regulatory role in the positive reinforcing properties of alcohol. In support of this hypothesis, several studies have also shown the inhibition of voluntary alcohol intake in CB1R knockout mice. It has been shown that young CB1R wild‐type mice exhibited a significantly higher alcohol preference and voluntary alcohol intake compared to their CB1 knockout littermates (Wang et al., 2003). Further, SR141716A has been shown to reduce voluntary alcohol intake in CB1R wild type but not in knockout mice (Wang et al., 2003). These observations clearly support the original hypothesis that inactivation of the CB1R contributes to the inhibition of alcohol intake in rodents. Similarly, administration of SR141716A significantly reduced alcohol and sucrose intake in C57BL/ 6x129/Ola mice and had no effect in CB1R knockout C57BL/6x129/Ola mice (Poncelet et al., 2003). Alcohol and sucrose intake were significantly reduced in CB1R knockout C57BL/6x129/Ola mice (Poncelet et al., 2003). A recent study also provides unequivocal evidence for the participation of the CB1R in the regulation of voluntary alcohol consumption and some of the acute intoxicating effects caused by administration of alcohol (Naassila et al., 2004). Alcohol consumption and preference are decreased, whereas alcohol sensitivity and withdrawal severity are increased, in CB1R knockout mice. The increased alcohol sensitivity is not related to low alcohol metabolism, because plasma alcohol concentrations after alcohol administration did not differ with either the genotype or the gender (Naassila et al., 2004). CB1R knockout mice showed an increase in alcohol withdrawal–induced convulsions, suggesting that alcohol consumption is also inversely related to alcohol withdrawal severity. Alcohol produced a similar reduction in body temperature in CB1 knockout and wild‐type mice (Racz et al., 2003). The motor coordination on rotarod was reduced in both CB1 knockout and wild‐type mice (Racz et al., 2003). In another study, CB1 knockout mice (CD1 background) were more sensitive to the hypothermic and sedative/hypnotic effects of alcohol than wild‐type mice (Naassila et al., 2004). CB1 knockout mice displayed a significant decrease in locomotor activity following injection of alcohol (1–2.5 g/kg) (Naassila et al., 2004). Alcohol self‐administrations and alcohol‐withdrawal severity have been shown to have an inverse genetic relationship (Metten et al., 1998). Rodents selectively bred for low alcohol intake showed greater alcohol withdrawal severity than did rodents selectively bred for high alcohol intake (Kosobud et al., 1988; Crabbe et al., 1994; Chester et al., 1998, 2003). Importantly, analysis of recombinant inbred strains for alcohol withdrawal severity led to identification of a quantitative trait locus on chromosome 4 in close proximity to the CB1R gene (Buck et al., 1997; Buck, 1998). Furthermore, CB1R expression has been shown to be increased in the rat medial frontal cortex after intermittent exposure to alcohol (Rimondini et al., 2002). However, chronic alcohol exposure has been shown to reduce CB1R gene expression in the rat Cpu, VMN, and CA1 and CA2 fields of the hippocampus and increase in the dentate gyrus (Ortiz et al., 2004a). Alcohol‐withdrawal symptoms observed in CB1R wild‐type mice were abolished in CB1R knockout mice (Racz et al., 2003). These data taken together indicate that the CB1R system could be important for alcohol‐reinforcing effects. These findings are significant for the development of potential therapeutic strategies for the treatment of alcoholism and addiction in general.

3

Therapeutic Opportunity

Although the detailed biochemistry, physiology, and pathophysiology of the endogenous cannabinoid system have not been fully determined, there is already overwhelming evidence indicating that a pharmacological modulation of the endogenous cannabinoid system could provide new tools for a number of disease states including drug addiction. > Table 18‐1 shows some potential therapeutic applications of CB1R agonists and antagonists. Excessive stimulation of the receptor leads to receptor toleration and is a particular problem of strong agonism (Kumar et al., 2001; Sim‐Selley and Martin, 2002). Several well‐ recognized side effects (sedation, cognitive dysfunction, tachycardia, postural hypotension, dry mouth, ataxia and immunosuppressant effects, and psychotropic effects) of cannabinoid agonists have hampered the use of these compounds in treatment protocols (for review, see Porter and Felder, 2001). Therefore, the

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. Table 18‐1 Potential therapeutic applications of cannabinoid drugs Cannabinoid drugs CB1 receptor agonists

CB1 receptor antagonists

Therapeutic application Treatment of cancer/postoperative pain Anticonvulsivant Antipastic in multiple sclerosis or spinal injury Treatment of memory deficits Treatment of obesity Treatment of alcohol dependence

For more details see following references (Nocerino et al., 2000; Hollister, 2001; Porter and Felder, 2001; Drysdale and Platt, 2003)

development of clinically acceptable weak agonists may be preferable for chronic use of cannabinoid‐based drugs to prevent receptor desensitization and also increase the therapeutic window. During disease, there are changes in endocannabinoid concentrations at the site of pathology (Baker et al., 2001; Basavarajappa and Hungund, 2002; Schabitz et al., 2002; Walker and Huang, 2002). Therefore, drugs that increase the level of endogenous cannabinoids by inhibiting their metabolism (FAAH inhibitors) or uptake could locally target sites of damage while limiting effects in uninvolved cognitive areas, and will thus have more selective therapeutic value. Cannabis is often used as a substitute for other, more dangerous drugs, including prescription narcotics, opiates, and alcohol. Cannabis has been proposed as a treatment for alcoholism (Mikuriya, 1970). However, a single controlled study of cannabis to treat alcoholics proved unsuccessful (Rosenberg et al., 1978; Rosenberg, 1979). Cannabinoid interactions with the dopamine system have been offered as a possible mechanism for some of the therapeutic potential of cannabinoid‐based drugs in alcoholism.

4

Conclusions

Much progress has been achieved in endocannabinoid system research within the last decade. Research on the molecular and neurobiological bases of the physiological/neurobehavioral role of the endocannabinoid system was hampered by the lack of specific research tools and technology. The situation started to change with the availability of molecular probes and other recombinant techniques that have led to major advances. Thus, we can look forward to a bright future of discoveries defining not only the role of the endocannabinoid system in normal brain function, but also on disease mechanisms that are poorly understood, like alcohol dependence, schizophrenia, anxiety, and other brain disorders. Future research should focus on (a) the more complete understanding of the mode of action of known endocannabinoids and cannabinoid receptors in the brain; (b) the molecular, physiological, and pharmacological characterization of missing key intermediates or elements of the endocannabinoid system, such as novel endocannabinoids and cannabinoid receptors in the brain; (c) their precise cellular and subcellular localization; (d) the biochemical pathways involved in endocannabinoid synthesis, uptake, and degradation; (e) the role played by the endocannabinoid system in modulating the release of neurotransmitters and other chemical messengers in various neurological and psychiatric disorders. Although these tasks are far from trivial, what is known so for about the endocannabinoid system suggests that they are well‐worth future research.

Acknowledgment The authors acknowledge support in part by grants from NIH AA11031 (BSB) and AA13003 (BLH). We thank Dr. Emmanuel S Onaivi for providing permission to reproduce CB1 gene structure.

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References Abel EL. 1970. Marijuana and memory. Nature 227: 1151-1152. Abood ME, Ditto KE, Noel MA, Showalter VM, Tao Q. 1997. Isolation and expression of a mouse CB1 cannabinoid receptor gene. Comparison of binding properties with those of native CB1 receptors in mouse brain and N18TG2 neuroblastoma cells. Biochem Pharmacol 53: 207-214. Adams IB, Compton DR, Martin BR. 1998. Assessment of anandamide interaction with the cannabinoid brain receptor: SR 141716A antagonism studies in mice and autoradiographic analysis of receptor binding in rat brain. J Pharmacol Exp Ther 284: 1209-1217. Akinshola BE, Chakrabarti A, Onaivi ES. 1999a. In‐vitro and in‐vivo action of cannabinoids. Neurochem Res 24: 1233-1240. Akinshola BE, Taylor RE, Ogunseitan AB, Onaivi ES. 1999b. Anandamide inhibition of recombinant AMPA receptor subunits in xenopus oocytes is increased by forskolin and 8‐bromo‐cyclic AMP. Naunyn Schmiedebergs Arch Pharmacol 360: 242-248. Aldrich MA. 1971.A brief legal history of marijuana. Presented to the Western Institute of Drug Problems Marijuana Conference, Portland Oregon, August 7, 14. Alger BE. 2002. Retrograde signaling in the regulation of synaptic transmission: Focus on endocannabinoids. Prog Neurobiol 68: 247-286. Alger BE, Pitler TA, Wagner JJ, Martin LA, Morishita W, et al. 1996. Retrograde signalling in depolarization‐induced suppression of inhibition in rat hippocampal CA1 cells. J Physiol 496: 197-209. Ameri A. 1999. The effects of cannabinoids on the brain. Prog Neurobiol 58: 315-348. Anwyl R. 1999. Metabotropic glutamate receptors: Electrophysiological properties and role in plasticity. Brain Res Rev 29: 83-120. Arnold JC, Topple AN, Mallet PE, Hunt GE, McGregor IS. 2001. The distribution of cannabinoid‐induced Fos expression in rat brain: Differences between the Lewis and Wistar strain. Brain Res 921: 240-255. Arnone M, Maruani J, Chaperon F, Thiebot M, Poncelet M, et al. 1997. Selective inhibition of sucrose and ethanol intake by SR 141716, an antagonist of central cannabinoid (CB1) receptors. Psychopharmacology 132: 104-106. Auclair N, Otani S, Soubrie P, Crepel F. 2000. Cannabinoids modulate synaptic strength and plasticity at glutamatergic synapses of rat prefrontal cortex pyramidal neurons. J Neurophysiol 83: 3287-3293. Baker D, Pryce G, Croxford JL, Brown P, Pertwee RG, et al. 2001. Endocannabinoids control spasticity in a multiple sclerosis model. FASEB J 15: 300-302.

Basavarajappa BS. 2005. Endocannabinoid system in the development of tolerance to alcohol. Klinik and Forschung (J Clin Res) 11: 16-19. Basavarajappa BS. 2006. Endocannabinoid signaling and alcohol addiction. In ‘‘New Research on Alcoholism’’ Douglas Baye, editor. New York: Nova Science Publishers, Inc USA (In press). Basavarajappa BS, Hungund BL. 1999a. Chronic Ethanol Increases the Cannabinoid Receptor Agonist, Anandamide and its Precursor N‐Arachidonylphosphatidyl ethanolamine in SK‐N‐SH Cells. J Neurochem 72: 522-528. Basavarajappa BS, Hungund BL. 1999b. Down‐regulation of cannabinoid receptor agonist‐stimulated [35S] GTPgS binding in synaptic plasma membrane from chronic ethanol exposed mouse. Brain Res 815: 89-97. Basavarajappa BS, Hungund BL. 2001. Cannabinoid receptor agonist‐stimulated [35S] guanosine triphosphategS binding in the brain of C57BL/6 and DBA/2 mice. J Neurosci Res 64: 429-436. Basavarajappa BS, Hungund BL. 2002. Neuromodulatory role of the endocannabinoid signaling system in alcoholism: An overview. Prostaglandins Leukot Essent Fatty Acids 66: 287-299. Basavarajappa BS, Hungund BL. 2005. Role of the endocannabinoid system in the development of tolerance to alcohol. Alcohol 40: 15-24. Basavarajappa BS, Cooper TB, Hungund BL. 1998. Chronic ethanol administration down‐regulates cannabinoid receptors in mouse brain synaptic plasma membrane. Brain Res 793: 212-218. Basavarajappa BS, Saito M, Cooper TB, Hungund BL. 2000. Stimulation of cannabinoid receptor agonist 2‐arachidonylglycerol by chronic ethanol and its modulation by specific neuromodulators in cerebellar granule neurons. Biochim Biophys Acta 1535: 78-86. Basavarajappa BS, Saito M, Cooper TB, Hungund BL. 2003. Chronic ethanol inhibits the anandamide transport and increases extracellular anandamide levels in cerebellar granule neurons. Eur J Pharmacol 466: 73-83. Beltramo M, Piomelli D. 2000. Carrier‐mediated transport and enzymatic hydrolysis of the endogenous cannabinoid 2‐arachidonylglycerol. Neuroreport 11: 1231-1235. Beltramo M, Stella N, Calignano A, Lin SY, Makriyannis A, et al. 1997. Functional role of high‐affinity anandamide transport, as revealed by selective inhibition. Science 277: 1094-1097. Ben‐Shabat S, Fride E, Sheskin T, Tamiri T, Rhee MH, et al. 1998. An entourage effect: Inactive endogenous fatty acid glycerol esters enhance 2‐arachidonoyl‐glycerol cannabinoid activity. Eur J Pharmacol 353: 23-31.

The endocannabinoid system Biegon A, Kerman IA. 2001. Autoradiographic study of pre‐ and postnatal distribution of cannabinoid receptors in human brain. NeuroImage 14: 1463-1468. Bisogno T, Sepe N, De Petrocellis L, Di Marzo V. 1997a. Biosynthesis of 2‐arachidonoyl‐glycerol, a novel cannabimimetic eicosanoid, in mouse neuroblastoma cells. Adv Exp Med Biol 433: 201-204. Bisogno T, Berrendero F, Ambrosino G, Cebeira M, Ramos JA, et al. 1999. Brain regional distribution of endocannabinoids: Implications for their biosynthesis and biological function. Biochem Biophys Res Commun 256: 377-380. Bisogno T, MacCarrone M, De Petrocellis L, Jarrahian A, Finazzi‐Agro A, et al. 2001. The uptake by cells of 2‐arachidonoylglycerol, an endogenous agonist of cannabinoid receptors. Biochem Biophys Res Commun 256: 377-380. Bisogno T, Sepe N, Melck D, Maurelli S, De Petrocellis L, et al. 1997b. Biosynthesis, release and degradation of the novel endogenous cannabimimetic metabolite 2‐arachidonoylglycerol in mouse neuroblastoma cells. Biochem J 322: 671-677. Bohme GA, Laville M, Ledent C, Parmentier M, Imperato A. 2000. Enhanced long‐term potentiation in mice lacking cannabinoid CB1 receptors. Neuroscience 95: 5-7. Bouaboula M, Bianchini L, McKenzie FR, Pouyssegur J, Casellas P. 1999b. Cannabinoid receptor CB1 activates the Na+/ H+ exchanger NHE‐1 isoform via Gi‐mediated mitogen activated protein kinase signaling transduction pathways. FEBS Lett 449: 61-65. Bouaboula M, Desnoyer N, Carayon P, Combes T, Casellas P. 1999a. Gi protein modulation induced by a selective inverse agonist for the peripheral cannabinoid receptor CB2: Implication for intracellular signalization cross‐regulation. Mol Pharmacol 55: 473-480. Bouaboula M, Bourrie B, Rinaldi‐Carmona M, Shire D, Le Fur G, et al. 1995a. Stimulation of cannabinoid receptor CB1 induces krox‐24 expression in human astrocytoma cells. J Biol Chem 270: 13973-13980. Bouaboula M, Perrachon S, Milligan L, Canat X, Rinaldi‐ Carmona M, et al. 1997. A selective inverse agonist for central cannabinoid receptor inhibits mitogen‐activated protein kinase activation stimulated by insulin or insulin‐ like growth factor 1. Evidence for a new model of receptor/ ligand interactions. J Biol Chem 272: 22330-22339. Bouaboula M, Poinot‐Chazel C, Bourrie B, Canat X, Calandra B, et al. 1995b. Activation of mitogen‐activated protein kinases by stimulation of the central cannabinoid receptor CBI. Biochem J 312: 637-641. Bracey MH, Hanson MA, Masuda KR, Stevens RC, Cravatt BF, 2002. Structural adaptations in a membrane enzyme that terminates endocannabinoid signaling. Science 298: 1793-1796.

14

Breivogel CS, Sim LJ, Childers SR. 1997. Regional differences in cannabinoid receptor/G‐protein coupling in rat brain. J Pharmacol Exp Ther 282: 1632-1642. Breivogel CS, Griffin G, Di Marzo V, Martin BR. 2001. Evidence for a new G protein‐coupled cannabinoid receptor in mouse brain. Mol Pharmacol 60: 155-163. Brocher S, Artola A, Singer W. 1992. Agonists of cholinergic and noradrenergic receptors facilitate synergistically the induction of long‐term potentiation in slices of rat visual cortex. Brain Res 573: 27-36. Bruinsma K, Taren DL. 1999. Chocolate: Food or drug? J Am Diet Assoc 99: 1249-1256. Buck KJ. 1998. Recent progress toward the identification of genes related to risk for alcoholism. Mamm Genome 9: 927-928. Buck KJ, Metten P, Belknap JK, Crabbe JC. 1997. Quantitative trait loci involved in genetic predisposition to acute alcohol withdrawal in mice. J Neurosci 17: 3946-3955. Buckley NE, McCoy KL, Mezey E, Bonner T, Zimmer A, et al. 2000. Immunomodulation by cannabinoids is absent in mice deficient for the cannabinoid CB(2) receptor. Eur J Pharmacol 396: 141-149. Buckley NE, Mezey E, Bonner T, Zimmer A, Felder CC, et al. 1997. Development of a CB2 knockout mouse. In: Proceedings of the Symposium on the Cannabinoids. Vermont, Berlington: International Cannabinoid Research Society. Cabral GA, Dove Pettit DA. 1998. Drugs and immunity: Cannabinoids and their role in decreased resistance to infectious disease. J Neuroimmunol 83: 116-123. Cadas H, di Tomaso E, Piomelli D. 1997. Occurrence and biosynthesis of endogenous cannabinoid precursor, N‐arachidonoyl phosphatidylethanolamine, in rat brain. J Neurosci 17: 1226-1242. Caenazzo MR, Hoehe W‐T, Hsieh WH, Berrettini TI, Bonner. et al. 1991. HindIII identifies a two allele DNA polymorphism of the human cannabinoid receptor gene (CNR). Nucleic Acids Res 19: 4798. Calandra B, Portier M, Kerneis A, Delpech M, Carillon C, et al. 1999. Dual intracellular signaling pathways mediated by the human cannabinoid CB1 receptor. Eur J Pharmacol 374: 445-455. Cao Q, Martinez M, Zhang J, Sanders AR, Badner JA, et al. 1997. Suggestive evidence for a schizophrenia susceptibility locus on chromosome 6q and a confirmation in an independent series of pedigrees. Genomics 43: 1-8. Cartmell J, Schoepp DD. 2000. Regulation of neurotransmitter release by metabotropic glutamate receptors. J Neurochem 75: 889-907. Caulfield MP, Brown DA. 1992. Cannabinoid receptor agonists inhibit Ca current in NG108‐15 neuroblastoma cells via a pertussis toxin‐sensitive mechanism. Br J Pharmacol 106: 231-232.

373

374

14

The endocannabinoid system

Caulfield MP, Robbins J, Brown DA. 1992. Neurotransmitters inhibit the omega‐conotoxin‐sensitive component of Ca current in neuroblastoma x glioma hybrid (NG 108‐15) cells, not the nifedipine‐sensitive component. Pflugers Arch 420: 486-492. Chakrabarti A, Onaivi ES, Chaudhuri G. 1995. Cloning and sequencing of a cDNA encoding the mouse brain‐type cannabinoid receptor protein. DNA Seq 6: 385-388. Chaperon F, Thiebot MH. 1999. Behavioral effects of cannabinoid agents in animals. Crit Rev Neurobiol 13: 243-281. Chester JA, Blose AM, Froehlich JC. 2003. Further evidence of an inverse genetic relationship between innate differences in alcohol preference and alcohol withdrawal magnitude in multiple selectively bred rat lines. Alcohol Clin Exp Res 27: 377-387. Chester JA, Risinger FO, Cunningham CL. 1998. Ethanol reward and aversion in mice bred for sensitivity to ethanol withdrawal. Alcohol Clin Exp Res 22: 468-473. Childers SR, Deadwyler SA. 1996. Role of cyclic AMP in the actions of cannabinoid receptors. Biochem Pharmacol 52: 819-827. Childers SR, Sexton T, Roy MB. 1994. Effects of anandamide on cannabinoid receptors in rat brain membranes. Biochem Pharmacol 47: 711-715. Christison R. 1848. in A Dispensatory or Commentary on the Pharmacopoeias of Great Britain (and the United States). Philadelphia: Lea and Blanchard; pp 971-974. Clement AB, Hawkins EG, Lichtman AH, Cravatt BF. 2003. Increased seizure susceptibility and proconvulsant activity of anandamide in mice lacking fatty acid amide hydrolase. J Neurosci 23: 3916-3923. Cohen C, Perrault G, Voltz C, Steinberg R, Soubrie P. 2002. SR141716, a central cannabinoid (CB1) receptor antagonist, blocks the motivational and dopamine‐releasing effects of nicotine in rats. Behav Pharmacol 13: 451-463. Colombo G, Agabio R, Fa M, Guano L, Lobina C, et al. 1998. Reduction of voluntary ethanol intake in ethanol‐preferring sP rats by the cannabinoid antagonist SR‐141716. Alcohol 33: 126-130. Colombo G, Serra S, Brunetti G, Gomez R, Melis S, et al. 2002. Stimulation of voluntary ethanol intake by cannabinoid receptor agonists in ethanol‐preferring sP rats. Psychopharmacology (Berl) 159: 181-187. Comings DE, Muhleman D, Gade R, Johnson JP, Verde R, et al. 1997. Cannabinoid receptor gene (CNR1): Association with i.v. drug use. Mol Psychiatry 2: 161-168. Conn PJ, Pin JP. 1997. Pharmacology and functions of metabotropic glutamate receptors. Annu Rev Pharmacol Toxicol 37: 205-237. Crabbe JC, Gallaher ES, Phillips TJ, Belknap JK. 1994. Genetic determinants of sensitivity to ethanol in inbred mice. Behav Neurosci 108: 186-195.

Cravatt BF, Demarest K, Patricelli MP, Bracey MH, Giang DK, et al. 2001. Supersensitivity to anandamide and enhanced endogenous cannabinoid signaling in mice lacking fatty acid amide hydrolase. Proc Natl Acad Sci USA 98: 9371-9376. Cravatt BF, Giang DK, Mayfield SP, Boger DL, Lerner RA, et al. 1996. Molecular characterization of an enzyme that degrades neuromodulatory fatty‐acid amides. Nature 384: 83-87. Cravatt BF, Prospero‐Garcia O, Siuzdak G, Gilula NB, Henriksen SJ, et al. 1995. Chemical characterization of a family of brain lipids that induce sleep. Science 268: 1506-1509. Dawson E. 1995. Identification of a polymorphic triplet marker for the brain cannabinoid receptor gene: Use in linkage and association studies of schizophrenia. Psychiatr Genet 5: S50. Day TA, Rakhshan F, Deutsch DG, Barker EL. 2001. Role of fatty acid amide hydrolase in the transport of the endogenous cannabinoid anandamide. Mol Pharmacol 59: 1369-1375. De Petrocellis L, Melck D, Bisogno T, Milone A, Di Marzo V. 1999. Finding of the endocannabinoid signalling system in Hydra, a very primitive organism: Possible role in the feeding response. Neuroscience 92: 377-387. De Petrocellis L, Melck D, Palmisano A, Bisogno T, Laezza C, et al. 1998. The endogeneous cannabinoid anandamide inhibits human breast cancer cell proliferation. Proc Natl Acad Sci USA 95: 8375-8380. Dean B, Sundram S, Bradbury R, Scarr E, Copolov D. 2001. Studies on [3H]CP‐55940 binding in the human central nervous system: Regional specific changes in density of cannabinoid‐1 receptors associated with schizophrenia and cannabis use. Neuroscience 103: 9-15. Denovan‐Wright EM, Robertson HA. 2000. Cannabinoid receptor messenger RNA levels decrease in a subset of neurons of the lateral striatum, cortex and hippocampus of transgenic Huntington’s disease mice. Neuroscience 98: 705-713. Derkinderen P, Toutant M, Burgaya F, Le Bert M, Siciliano JC, et al. 1996. Regulation of a neuronal form of focal adhesion kinase by anandamide. Science 273: 1719-1722. Derkinderen P, Toutant M, Kadare G, Ledent C, Parmentier M, et al. 2001. Dual role of Fyn in the regulation of FAK+6,7 by cannabinoids in hippocampus. J Biol Chem 276: 38289-38296. Deutsch DG, Chin SA. 1993. Enzymatic synthesis and degradation of anandamide, a cannabinoid receptor agonist. Biochem Pharmacol 46: 791-796. Deutsch DG, Glaser ST, Howell JM, Kunz JS, Puffenbarger RA, et al. 2001. The cellular uptake of anandamide is coupled to its breakdown by fatty‐acid amide hydrolase. J Biol Chem 276: 6967-6973.

The endocannabinoid system Devane WA, Dysarz FAI, Johnson MR, Melvin LS, Howlett AC. 1988. Determination and characterization of a cannabinoid receptor in rat brain. Mol Pharmacol 34: 605-613. Devane WA, Hanus L, Breuer A, Pertwee RG, Stevenson LA, et al. 1992. Isolation and structure of a brain constituent that binds to the cannabinoid receptor. Science 258: 1946-1949. Dewey WL. 1986. Cannabinoid pharmacology. Pharmacol Rev 38: 151-178. Di Chiara G, Imperato A. 1988. Drugs abused by humans preferentially increase synaptic dopamine concentrations in the mesolimbic system of freely moving rats. Proc Natl Acad Sci USA 85: 5274-5278. Di Marzo V. 1998. ‘Endocannabinoids’ and other fatty acid derivatives with cannabimimetic properties: biochemistry and possible. Biochim Biophys Acta 1392: 153-175. Di Marzo V, Bisogno T, De Petrocellis L. 2000a. Endocannabinoids: New targets for drug development. Curr Pharm Des 6: 1361-1380. Di Marzo V, De Petrocellis L, Bisogno T, Melck D. 1999. Metabolism of anandamide and 2‐arachidonoylglycerol: An historical overview and some recent developments. Lipids 34(Suppl): S319-325. Di Marzo V, De Petrocellis L, Sepe N, Buono A. 1996. Biosynthesis of anandamide and related acylethanolamides in mouse J774 macrophages and N18 neuroblastoma cells. Biochem J 316: 977-984. Di Marzo V, Bisogno T, Sugiura T, Melck D, De Petrocellis L. 1998a. The novel endogenous cannabinoid 2‐arachidonoylglycerol is inactivated by neuronal‐ and basophil‐like cells: Connections with anandamide. Biochem J 331: 15-19. Di Marzo V, Hill MP, Bisogno T, Crossman AR, Brotchie JM. 2000b. Enhanced levels of endogenous cannabinoids in the globus pallidus are associated with a reduction in movement in an animal model of Parkinson’s disease. FASEB J 14: 1432-1438. Di Marzo V, Bisogno T, Melck D, Ross R, Brockie H, et al. 1998c. Interactions between synthetic vanilloids and the endogenous cannabinoid system. FEBS Lett 436: 449-454. Di Marzo V, Breivogel CS, Tao Q, Bridgen DT, Razdan RK, et al. 2000c. Levels, metabolism, and pharmacological activity of anandamide in CB(1) cannabinoid receptor knockout mice: Evidence for non‐CB(1), non‐CB(2) receptor‐mediated actions of anandamide in mouse brain. J Neurochem 75: 2434-2444. Di Marzo V, Fontana A, Cadas H, Schinelli S, Cimino G, et al. 1994. Formation and inactivation of endogenous cannabinoid anandamide in central neurons. Nature 372: 686-691. Di Marzo V, Sepe N, De Petrocellis L, Berger A, Crozier G, et al. 1998b. Trick or treat from food endocannabinoids? Nature 396: 636-637.

14

di Tomaso E, Beltramo M, Piomelli D. 1996. Brain cannabinoids in chocolate. Nature 382: 677-678. Diana MA, Levenes C, Mackie K, Marty A. 2002. Short‐term retrograde inhibition of GABAergic synaptic currents in rat Purkinje cells is mediated by endogenous cannabinoids. J Neurosci 22: 200-208. Dinh TP, Carpenter D, Leslie FM, Freund TF, Katona I, et al. 2002. Brain monoglyceride lipase participating in endocannabinoid inactivation. Proc Natl Acad Sci USA 99: 10819-10824. Doherty J, Dingledine R. 2003. Functional interactions between cannabinoid and metabotropic glutamate receptors in the central nervous system. Curr Opin Pharmacol 3: 46-53. Drysdale AJ, Platt B. 2003. Cannabinoids: Mechanisms and therapeutic applications in the CNS. Curr Med Chem 10: 2719-2732. Egertova M, Elphick MR. 2000. Localisation of cannabinoid receptors in the rat brain using antibodies to the intracellular C‐terminal tail of CB. J Comp Neurol 422: 159-171. Egertova M, Cravatt BF, Elphick MR. 2003. Comparative analysis of fatty acid amide hydrolase and cb(1) cannabinoid receptor expression in the mouse brain: Evidence of a widespread role for fatty acid amide hydrolase in regulation of endocannabinoid signaling. Neuroscience 119: 481-496. Egertova M, Giang DK, Cravatt BF, Elphick MR. 1998. A new perspective on cannabinoid signalling: Complementary localization of fatty acid amide hydrolase and the CB1 receptor in rat brain. Proc R Soc Lond B Biol Sci 265: 2081-2085. Facci L, Dal Toso R, Romanello S, Buriani A, Skaper SD, et al. 1995. Mast cells express a peripheral cannabinoid receptor with differential sensitivity to anandamide and palmitoylethanolamide. Proc Natl Acad Sci USA 92: 3376-3380. Farooqui AA, Taylor WA, Horrocks LA. 1986. Characterization and solubilization of membrane bound diacylglycerol lipases from bovine brain. Int J Biochem 18: 991-997. Felder C, Nielsen A, Briley E, Palkovits M, Priller J, et al. 1996. Isolation and measurement of the endogenous cannabinoid receptor agonist, anandamide, in brain and peripheral tissues of human and rat. FEBS Lett 393: 231-235. Felder CC, Briley EM, Axelrod J, Simpson JT, Mackie K, et al. 1993. Anandamide, an endogenous cannabimimetic eicosanoid, binds to the cloned human cannabinoid receptor and stimulates receptor‐mediated signal transduction. Proc Natl Acad Sci USA 90: 7656-7660. Felder CC, Joyce KE, Briley EM, Mansouri J, Mackie K, et al. 1995. Comparison of the pharmacology and signal transduction of the human cannabinoid CB1 and CB2 receptors. Mol Pharmacol 48: 443-450. Ferrer B, Asbrock N, Kathuria S, Piomelli D, Giuffrida A. 2003. Effects of levodopa on endocannabinoid levels

375

376

14

The endocannabinoid system

in rat basal ganglia: Implications for the treatment of levodopa‐induced dyskinesias. Eur J Neurosci 18: 1607-1614. Fowler CJ, Nilsson O, Andersson M, Disney G, Jacobsson SO, et al. 2001. Pharmacological properties of cannabinoid receptors in the avian brain: Similarity of fat and chicken cannabinoidl receptor recognition sites and expression of cannabinoid2 receptor‐like immunoreactivity in the embryonic chick brain. Pharmacol Toxicol 88: 213-222. Frazier J. 1974. The Marijuana Farmers, Hemp Cults and Cultures. New Orleans, LA: Solar Age Press; p. 133. Freedland CS, Sharpe AL, Samson HH, Porrino LJ. 2001. Effects of SR141716A on ethanol and sucrose self‐ administration. Alcohol Clin Exp Res 25: 277-282. Freund TF, Katona I, Piomelli D. 2003. Role of endogenous cannabinoids in synaptic signaling. Physiol Rev 83: 1017-1066. Fritzsche M. 2000. Are cannabinoid receptor knockout mice animal models of schizophrenia. Med Hypotheses 56: 638-643. Gadzicki D, Muller‐Vahl K, Stuhrmann M. 1999. A frequent polymorphism in the coding exon of the human cannabinoid receptor (CNR1) gene. Mol Cell Probes 13: 321-323. Gallate JE, McGregor IS. 1999. The motivation for beer in rats: Effects of ritanserin, naloxone and SR 141716. Psychopharmacology 142: 302-308. Gallate JE, Saharov T, Mallet PE, McGregor IS. 1999. Increased motivation for beer in rats following administration of a cannabinoid CB1 receptor agonist. Eur J Pharmacol 370: 233-240. Gaoni Y, Mechoulam R. 1964. Isolation, structure and partial synthesis of an active constituent of hashish. J Am Chem Soc 86: 1646-1647. Gebremedhin D, Lange AR, Campbell WB, Hillard CJ, Harder DR. 1999. Cannabinoid CB1 receptor of cat cerebral arterial muscle functions to inhibit L‐type Ca2+ channel current. Am J Physiol 276: H2085-2093. Gerard CM, Mollereau C, Vassart G, Parmentier M. 1991. Molecular cloning of a human cannabinoid receptor which is also expressed in testis. Biochem J 279: 129-134. Gerdeman G, Lovinger DM. 2001. CB1 cannabinoid receptor inhibits synaptic release of glutamate in rat dorsolateral striatum. J Neurophysiol 85: 468-471. Gerdeman GL, Ronesi J, Lovinger DM. 2002. Postsynaptic endocannabinoid release is critical to long‐term depression in the striatum. Nat Neurosci 5: 446-451. Gessa GL, Melis M, Muntoni AL, Diana M. 1998. Cannabinoids activate mesolimbic dopamine neurons by an action on cannabinoid CB1 receptors. Eur J Pharmacol 341: 39-44. Giuffrida A, Beltramo M, Piomelli D. 2001. Mechanisms of endocannabinoid inactivation: Biochemistry and pharmacology. J Pharmacol Exp Ther 298: 7-14.

Giuffrida A, Parsons LH, Kerr TM, Rodriguez de Fonseca F, et al. 1999. Dopamine activation of endogenous cannabinoid signaling in dorsal striatum. Nat Neurosci 2: 358-363. Glaser ST, Abumrad NA, Fatade F, Kaczocha M, Studholme KM, et al. 2003. Evidence against the presence of an anandamide transporter. Proc Natl Acad Sci USA 100: 4269-4274. Glass M. 2001. The role of cannabinoids in neurodegenerative diseases. Prog Neuropsychopharmacol Biol Psychiatry 25: 743-765. Glass M, Felder CC. 1997. Concurrent stimulation of cannabinoid CB1 and dopamine D2 receptors augments cAMP accumulation in striatal neurons: Evidence for a Gs linkage to the CB1 receptor. J Neurosci 17: 5327-5333. Glass M, Dragunow M, Faull RL. 1997. Cannabinoid receptors in the human brain: A detailed anatomical and quantitative autoradiographic study in the fetal, neonatal and adult human brain. Neuroscience 77: 299-318. Glass M, Dragunow M, Faull RL. 2000. The pattern of neurodegeneration in Huntington’s disease: A comparative study of cannabinoid, dopamine, adenosine and GABA(A) receptor alterations in the human basal ganglia in Huntington’s disease. Neuroscience 97: 505-519. Glass M, van Dellen A, Blakemore C, Hannan AJ, Faull RL. 2004. Delayed onset of huntington’s disease in mice in an enriched environment correlates with delayed loss of cannabinoid CB1 receptors. Neuroscience 123: 207-212. Godwin H. 1967. Pollen Analytic Evidence for the Cultivation of Cannabis in England. Rev Paleobotany Palnyol 4: 71-80. Gomez del Pulgar T, Velasco G, Guzman M. 2000. The CB1 cannabinoid receptor is coupled to the activation of protein kinase B/Akt. Biochem J 347: 369-373. Gonzalez S, Grazia Cascio M, Fernandez‐Ruiz J, Fezza F, Di Marzo V, et al. 2002. Changes in endocannabinoid contents in the brain of rats chronically exposed to nicotine, ethanol or cocaine. Brain Res 954: 73. Goparaju SK, Ueda N, Yamaguchi H, Yamamoto S. 1998. Anandamide amidohydrolase reacting with 2‐arachidonoylglycerol, another cannabinoid receptor ligand. FEBS Lett 422: 69-73. Griffin G, Tao Q, Abood ME. 2000. Cloning and pharmacological characterization of the rat CB(2) cannabinoid receptor. J Pharmacol Exp Ther 292: 886-894. Guzman M, Sanchez C. 1999. Effects of cannabinoids on energy metabolism. Life Sci 65: 657-664. Guzman M, Galve‐Roperh I, Sanchez C. 2001. Ceramide: A new second messenger of cannabinoid action. Trends Pharmacol Sci 22: 19-22. Hajos N, Ledent C, Freund TF. 2001. Novel cannabinoid‐ sensitive receptor mediates inhibition of glutamatergic synaptic transmission in the hippocampus. Neuroscience 106: 1-4.

The endocannabinoid system Hajos N, Katona I, Naiem SS, Mac Kie K, Ledent C, et al. 2000. Cannabinoids inhibit hippocampal GABAergic transmission and network oscillations. Eur J Neurosci 12: 3239-3249. Hampson RE, Mu J, Deadwyler SA. 2000. Cannabinoid and kappa opioid receptors reduce potassium K current via activation of G(s) proteins in cultured hippocampal neurons. J Neurophysiol 84: 2356-2364. Hansen HH, Hansen SH, Schousboe A, Hansen HS. 2000. Determination of the phospholipid precursor of anandamide and other N‐acylethanolamine phospholipids before and after sodium azide‐induced toxicity in cultured neocortical neurons. J Neurochem 75: 861-871. Hanus L, Gopher A, Almog S, Mechoulam R. 1993. Two new unsaturated fatty acid ethanolamides in brain that bind to the cannabinoid receptor. J Med Chem 36: 3032-3034. Hanus L, Abu‐Lafi S, Fride E, Breuer A, Vogel Z, et al. 2001. 2‐arachidonyl glyceryl ether, an endogenous agonist of the cannabinoid CB1 receptor. Proc Natl Acad Sci USA 98: 3662-3665. Herkenham M, Lynn AB, Little MD, Johnson MR, Melvin LS, et al. 1990. Cannabinoid receptor localization in brain. Proc Natl Acad Sci USA 87: 1932-1936. Herkenham M, Lynn AB, Johnson MR, Melvin LS, de Cost BR, et al. 1991. Characterization and localization of cannabinoid receptors in rat brain: A quantitative in vitro autoradiographic study. J Neurosci 16: 8057-8066. Hillard CJ, Jarrahian A. 2000. The movement of N‐arachidonoylethanolamine (anandamide) across cellular membranes. Chem Phys Lipids 108: 123-134. Hillard CJ, Jarrahian A. 2003. Cellular accumulation of anandamide: Consensus and controversy. Br J Pharmacol 140: 802-808. Hillard CJ, Edgemond WS, Jarrahian A, Campbell WB. 1997. Accumulation of N‐arachidonoylethanolamine (anandamide) into cerebellar granule cells occurs via facilitated diffusion. J Neurochem 69: 631-638. Ho BY, Uezono Y, Takada S, Takase I, Izumi F. 1999. Coupling of the expressed cannabinoid CB1 and CB2 receptors to phospholipase C and G protein‐coupled inwardly rectifying K+ channels. Receptors Channels 6: 363-374. Hoehe MR, Caenazzo L, Martinez MM, Hsieh WT, Modi WS, et al. 1991. Genetic and physical mapping of the human cannabinoid receptor gene to chromosome 6q14‐q15. New Biol 3: 880-885. Hoffman AF, Lupica CR. 2000. Mechanisms of cannabinoid inhibition of GABA(A) synaptic transmission in the hippocampus. J Neurosci 20: 2470-2479. Hollister LE. 1986. Health aspects of cannabis. Pharmacol Rev 38: 1-20. Hollister LE. 2001. Marijuana (Cannabis) as medicine. J Cnnabis Ther 1: 5-27.

14

Howlett AC, Mukhopadhyay S. 2000. Cellular signal transduction by anandamide and 2‐arachidonoylglycerol. Chem Phys Lipids 108: 53-70. Howlett AC, Johnson MR, Melvin LS, Milne GM. 1988. Nonclassical cannabinoid analgetics inhibit adenylate cyclase: Development of a cannabinoid receptor model. Mol Pharmacol 33: 297-302. Howlett AC, Barth F, Bonner TI, Cabral G, Casellas P, et al. 2002. International Union of Pharmacology. XXVII. Classification of cannabinoid receptors. Pharmacol Rev 54: 161-202. Huang CC, Lo SW, Hsu KS. 2001. Presynaptic mechanisms underlying cannabinoid inhibition of excitatory synaptic transmission in rat striatal neurons. J Physiol 532(Pt3): 731-748. Huang SM, Bisogno T, Trevisani M, Al‐Hayani A, De Petrocellis L, et al. 2002. An endogenous capsaicin‐like substance with high potency at recombinant and native vanilloid VR1 receptors. Proc Natl Acad Sci USA 99: 8400-8405. Hungund BL, Basavarajappa BS. 2000a. Are anandamide and cannabinoid receptors involved in ethanol tolerance? A review of the evidence. Alcohol 35: 126-133. Hungund BL, Basavarajappa BS. 2000b. Role of brain’s own marijuana, anandamide and its cannabinoid receptors (CB1) in alcoholism. Recent Research Development in Neurochemistry. Vol.3 S.G. Pandalai R.S. editor. Research Signpost, India: Trivandrum; pp. 9–26. Hungund BL, Basavarajappa BS. 2000c. Distinct differences in the cannabinoid receptor binding in the brain of C57BL/6 and DBA/2 mice, selected for their differences in voluntary ethanol consumption. J Neurosci Res 60: 122-128. Hungund BL, Basavarajappa BS. 2004. Role of endocannabinoids and cannabinoid CB1 receptors in alcohol‐related behaviours. Ann N Y Acad Sci 1025: 1-14. Hungund BL, Szakall I, Adam A, Basavarajappa BS, Vadasz C. 2003. Cannabinoid CB1 receptor knockout mice exhibit markedly reduced voluntary alcohol consumption and lack alcohol‐induced dopamine release in the nucleus accumbens. J Neurochem 84: 698-704. Hungund BL, Basavarajappa BS, Vadasz C, Kunos G, Rodriguez de, et al. 2002. Ethanol, endocannabinoids and cannabinoidergic signaling system. Alcohol Clin Exp Res 26: 565-574. Hungund BL, Vinod KY, Kassir SA, Basavarajappa BS, Yalamanchili R, et al. (2004). Upregulation of CB1 receptors and agonist‐stimulated [35S]GTPgammaS binding in the prefrontal cortex of depressed suicide victims. Mol Psychiatry 9: 184-190. Jarai Z, Wagner JA, Varga K, Lake KD, Compton DR, et al. 1999. Cannabinoidinduced mesenteric vasodilation through

377

378

14

The endocannabinoid system

an endothelial site distinct from CB1 or CB2 receptors. Proc Natl Acad Sci USA 96: 14136-14141. Jarrahian A, Manna S, Edgemond W, Campbell W, Hillard C. 2000. Structure—activity relationships among N‐Arachidonylethanolamine (Anandamide) head group analogues for the anandamide transporter. J Neurochem 74: 2597-2606. Johnson JP, Muhleman D, MacMurray J, Gade R, Verde R, et al. 1997. Association between the cannabinoid receptor gene (CNR1) and the P300 event‐related potential. Mol Psychiatry 2: 169-171. Kabelik J, Krejci Z, Santavy F. 1960. Cannabis as a Medicant. Bull Narc 12: 5-23. Katayama K, Ueda N, Katoh I, Yamamoto S. 1999. Equilibrium in the hydrolysis and synthesis of cannabimimetic anandamide demonstrated by a purified enzyme. Biochim Biophys Acta 1440: 205-214. Kathmann M, Haug K, Heils A, Nothen MM, Schlicker E. 2000. Exchange of three amino acids in the cannabinoid CB1 receptor (CNR1) of an epilepsy patient. Proceedings of the Symposium on the Cannabinoids, Vermont, Burlington: ICRS; p. 18. Katona I, Rancz EA, Acsady L, Ledent C, Mackie K, et al. 2001. Distribution of CB1 cannabinoid receptors in the amygdala and their role in the control of GABAergic transmission. J Neurosci 21: 9506-9518. Katona I, Sperlagh B, Sik A, Kafalvi A, Vizi ES, et al. 1999. Presynaptically located CB1 cannabinoid receptors regulate GABA release from axon terminals of specific hippocampal interneurons. J Neurosci 19: 4544-4558. Katsuki H, Izumi Y, Zorumski CF. 1997. Noradrenergic regulation of synaptic plasticity in the hippocampal CA1 region. J Neurophysiol 77: 3013-3020. Kearn CS, Greenberg MJ, DiCamelli R, Kurzawa K, Hillard CJ. 1999. Relationships between ligand affinities for the cerebellar cannabinoid receptor CB1 and the induction of GDP/GTP exchange. J Neurochem 72: 2379-2387. Kim J, Isokawa M, Ledent C, Alger BE. 2002. Activation of muscarinic acetylcholine receptors enhances the release of endogenous cannabinoids in the hippocampus. J Neurosci 22: 10182-10191. Kobayashi M, Ohno M, Shibata S, Yamamoto T, Watanabe S. 1997. Concurrent blockade of beta‐adrenergic and muscarinic receptors suppresses synergistically long‐term potentiation of population spikes in the rat hippocampal CA1 region. Brain Res 777: 242-246. Kondo S, Kondo H, Nakane S, Kodaka T, Tokumura A, et al. 1998. 2‐Arachidonoylglycerol, an endogenous cannabinoid receptor agonist: Identification as one of the major species of monoacylglycerols in various rat tissues, and evidence for its generation through CA2+‐ dependent and ‐independent mechanisms. FEBS Lett 429: 152-156.

Konrad RJ, Major CD, Wolf BA. 1994. Diacylglycerol hydrolysis to arachidonic acid is necessary for insulin secretion from isolated pancreatic islets: Sequential actions of diacylglycerol and monoacylglycerol lipases. Biochemistry 33: 13284-13294. Kosobud A, Bodor AS, Crabbe JC. 1988. Voluntary consumption of ethanol in WSP, WSC and WSR selectively bred mouse lines. Pharmacol Biochem Behav 29: 601-607. Kreitzer AC, Regehr WG. 2001a. Cerebellar depolarization‐ induced suppression of inhibition is mediated by endogenous cannabinoids. J Neurosci 21: RC174. Kreitzer AC, Regehr WG. 2001b. Retrograde inhibition of presynaptic calcium influx by endogenous cannabinoids at excitatory synapses onto Purkinje cells. Neuron 29: 717-727. Kumar RN, Chambers WA, Pertwee RG. 2001. Pharmacological actions and therapeutic uses of cannabis and cannabinoids. Anaesthesia 56: 1059-1068. Kurahashi Y, Ueda N, Suzuki H, Suzuki M, Yamamoto S. 1997. Reversible hydrolysis and synthesis of anandamide demonstrated by recombinant rat fatty‐acid amide hydrolase. Biochem Biophys Res Commun 237: 512-515. Lallemand F, Soubrie PH, De Witte PH. 2001. Effects of CB1 cannabinoid receptor blockade on ethanol preference after chronic ethanol administration. Alcohol Clin Exp Res 25: 1317-1323. Ledent C, Valverde O, Cossu G, Petitet F, Aubert JF, et al. 1999. Unresponsiveness to cannabinoids and reduced addictive effects of opiates in CB1 receptor knockout mice. Science 283: 401-404. Leweke FM, Schneider U, Thies M, Munte TF, Emrich HM. 1999. Effects of synthetic Delta(9)‐tetrahydrocannabinol on binocular depth inversion of natural and artificial objects in man. Psychopharmacology 142: 230-235. Li T, Liu X, Zhu ZH, Zhao J, Hu X, et al. 2000. No association between (AAT)n repeats in the cannabinoid receptor gene (CNR1) and heroin abuse in a Chinese population. Mol Psychiatry 5: 128-130. Lichtman AH, Hawkins EG, Griffin G, Cravatt BF. (2002). Pharmacological activity of fatty acid amides is regulated, but not mediated, by fatty acid amide hydrolase in vivo. J Pharmacol Exp Ther 302: 73-79. Liu J, Gao B, Mirshahi F, Sanyal AJ, Khanolkar AD, et al. 2000. Functional CB1 cannabinoid receptors in human vascular endothelial cells. Biochem J Mar 15; 346 Pt 3: 835-40. Llano I, Leresche N, Marty A. 1991. Calcium entry increases the sensitivity of cerebellar Purkinje cells to applied GABA and decreases inhibitory synaptic currents. Neuron 6: 565-574. Loewe S. 1950. Cannabiswiswirkstoffe and Parmakologie der Cannabinole (Active constituent of cannabis and pharmacology of the cannabinoids.) (in German.). Arciv Fur Experimentelle pathologie und pharmakogie 175‐193.

The endocannabinoid system Maccarrone M, Bari M, Menichelli A, Del Principe D, Agro AF. 1999. Anandamide activates human platelets through a pathway independent of the arachidonate cascade. FEBS Lett 447: 277-282. Maccarrone M, Attina M, Bari M, Cartoni A, Ledent C, et al. 2001. Anandamide degradation and N‐acylethanolamines level in wild‐type and CB1 cannabinoid receptor knockout mice of different ages. J Neurochem 78: 339-348. Maccarrone M, Bari M, Lorenzon T, Bisogno T, Di Marzo V, 2000b. Anandamide uptake by human endothelial cells and its regulation by nitric oxide. J Biol Chem 275: 13484-13492. Maccarrone M, Fiorucci L, Erba F, Bari M, Finazzi‐Agro A, et al. 2000a. Human mast cells take up and hydrolyze anandamide under the control of 5‐lipoxygenase and do not express cannabinoid receptors. FEBS Lett 468: 176-180. Maccarrone M, van der Stelt M, Rossi A, Veldink GA, Vliegenthart JF, et al. 1998. Anandamide hydrolysis by human cells in culture and brain. J Biol Chem 273: 32332-32339. Mackie K, Hille B. 1992. Cannabinoids inhibit N‐type calcium channels in neuroblastoma‐glioma cells. Proc Natl Acad Sci USA 89: 3825-3829. Maejima T, Ohno‐Shosaku T, Kano M. 2001. Endogenous cannabinoid as a retrograde messenger from depolarized postsynaptic neurons to presynaptic terminals. Neurosci Res 40: 205-210. Mailleux P, Verslype M, Preud’homme X, Vanderhaeghen JJ. 1994. Activation of multiple transcription factor genes by tetrahydrocannabinol in rat forebrain. Neuroreport 5: 1265-1268. Maneuf YP, Nash JE, Crossman AR, Brotchie JM. 1996. Activation of the cannabinoid receptor by delta 9‐tetrahydrocannabinol reduces gamma‐aminobutyric acid uptake in the globus pallidus. Eur J Pharmacol 308: 161-164. Marsicano G, Wotjak CT, Azad SC, Bisogno T, Rammes G, et al. 2002. The endogenous cannabinoid system controls extinction of aversive memories. Nature 418: 530-534. Martinez M, Goldin LR, Cao Q, Zhang J, Sanders AR, et al. 1999. Follow‐up study on a susceptibility locus for schizophrenia on chromosome 6q. Am J Med Genet 88: 337-343. Mascia MS, Obinu MC, Ledent C, Parmentier M, Bohme GA, et al. 1999. Lack of morphine‐induced dopamine release in the nucleus accumbens of cannabinoid CB(1) receptor knockout mice. Eur J Pharmacol 383: R1-2. Mato S, Pazos A, Valdizan EM. 2002. Cannabinoid receptor antagonism and inverse agonism in response to SR141716A on cAMP production in human and rat brain. Eur J Pharmacol 443: 43-46. Matsuda LA. 1997. Molecular aspects of cannabinoid receptors. Crit Rev Neurobiol 11: 143-166. Matsuda LA, Bonner TI, Lolait SJ. 1993. Localization of cannabinoid receptor mRNA in rat brain. J Comp Neurol 327: 535-550.

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Matsuda LA, Lolait SJ, Brownstein MJ, Young AC, Bonner TI. 1990. Structure of a cannabinoid receptor and functional expression of the cloned cDNA. Nature 346: 561-564. Mazzola C, Micale V, Drago F. 2003. Amnesia induced by beta‐amyloid fragments is counteracted by cannabinoid CB1 receptor blockade. Eur J Pharmacol 477: 219-225. McGregor IS, Arnold JC, Weber MF, Topple AN, Hunt GE. 1998. A comparison of delta 9‐THC and anandamide induced c‐fos expression in the rat forebrain. Brain Res 802: 19-26. Mechoulam R, Fride E, Di Marzo V. 1998. Endocannabinoids. Eur J Pharmacol 359: 1-18. Mechoulam R, Ben‐Shabat S, Hanus L, Ligumsky M, Kaminski NE, et al. 1995. Identification of an endogenous 2‐monoglyceride, present in canine gut, that binds to cannabinoid receptors. Biochem Pharmacol 50: 83-90. Melck D, Bisogno T, De Petrocellis L, Chuang H, Julius D, et al. 1999. Unsaturated long‐chain N‐acyl‐vanillyl‐amides (N‐AVAMs): Vanilloid receptor ligands that inhibit anandamide‐facilitated transport and bind to CB1 cannabinoid receptors. Biochem Biophys Res Commun 262: 275-284. Melis M, Pistis M, Perra S, Muntoni AL, Pillolla G, et al. 2004. Endocannabinoids mediate presynaptic inhibition of glutamatergic transmission in rat ventral tegmental area dopamine neurons through activation of CB1 receptors. J Neurosci 24: 53-62. Meschler JP, Kraichely DM, Wilken GH, Howlett AC. 2000. Inverse agonist properties of N‐(piperidin‐1‐yl)‐5‐(4‐ chlorophenyl)‐1‐(2, 4‐dichlorophenyl)‐4‐methyl‐1H‐pyrazole‐3‐carboxamide HC1 (SR141716A) and 1‐(2‐chlorophenyl)‐4‐cyano‐5‐(4‐methoxyphenyl)‐1H‐pyrazole‐3‐carboxyl ic acid phenylamide (CP‐272871) for the CB(1) cannabinoid receptor. Biochem Pharmacol 60: 1315-1323. Metten P, Phillips TJ, Crabbe JC, Tarantino LM, McClearn GE, et al. 1998. High genetic susceptibility to ethanol withdrawal predicts low ethanol consumption. Mamm Genome 9: 983-990. Mikuriya TH. 1970. Cannabis substitution. An adjunctive therapeutic tool in the treatment of alcoholism. Med Times 98: 187-191. Morishita W, Alger BE. 2000. Differential effects of the group II mGluR agonist, DCG‐IV, on depolarization‐induced suppression of inhibition in hippocampal CA1 and CA3 neurons. Hippocampus 10: 261-268. Morishita W, Alger BE. 2001. Direct depolarization and antidromic action potentials transiently suppress dendritic IPSPs in hippocampal CA1 pyramidal cells. J Neurophysiol 85: 480-484. Morisset V, Urban L. 2001. Cannabinoid‐induced presynaptic inhibition of glutamatergic EPSCs in substantia gelatinosa neurons of the rat spinal cord. J Neurophysiol 86: 40-48.

379

380

14

The endocannabinoid system

Mu J, Zhuang SY, Kirby MT, Hampson RE, Deadwyler SA. 1999. Cannabinoid receptors differentially modulate potassium A and D currents in hippocampal neurons in culture. J Pharmacol Exp Ther 291: 893-902. Munro S, Thomas KL, Abu‐Shaar M. 1993. Molecular characterization of a peripheral receptor for cannabinoids. Nature 365: 61-65. Musty RE, Deyo RA, Baer JL, Darrow SM, Coleman B. 2000. Effects of SR141716 on animal models of schizophrenia. Symposium on the cannabinoids, International Cannabinoid Research Society. Vermont: Burlington. Naassila M, Pierrefiche O, Ledent C, Daoust M. 2004. Decreased alcohol self‐administration and increased alcohol sensitivity and withdrawal in CB1 receptor knockout mice. Neuropharmacology 46: 243-253. Natarajan V, Reddy PV, Schmid PC, Schmid HH. 1981. On the biosynthesis and metabolism of N‐acylethanolamine phospholipids in infarcted dog heart. Biochim Biophys Acta 664: 445-448. Netzeband JG, Conroy SM, Parsons KL, Gruol DL. 1999. Cannabinoids enhance NMDA‐elicited Ca2+ signals in cerebellar granule neurons in culture. J Neurosci 19: 8765-8777. Nocerino E, Amato M, Izzo AA. 2000. Cannabis and cannabinoid receptors. Fitoterapia 71: S6-12. Ohno‐Shosaku T, Maejima T, Kano M. 2001. Endogenous cannabinoids mediate retrograde signals from depolarized postsynaptic neurons to presynaptic terminal. Neuron 29: 729-738. Ohno‐Shosaku T, Sawada S, Kano M. 2000. Heterosynaptic expression of depolarization‐induced suppression of inhibition (DSI) in rat hippocampal cultures. Neurosci Res 36: 67-71. Ohno‐Shosaku T, Sawada S, Yamamoto C. 1998. Properties of depolarization‐induced suppression of inhibitory transmission in cultured rat hippocampal neurons. Pflugers Arch 435: 273-279. Ohno‐Shosaku T, Tsubokawa H, Mizushima I, Yoneda N, Zimmer A, et al. 2002. Presynaptic cannabinoid sensitivity is a major determinant of depolarization‐induced retrograde suppression at hippocampal synapses. J Neurosci 22: 3864-3872. Okamoto Y, Morishita J, Tsuboi K, Tonai T, Ueda N. 2004. Molecular characterization of a phospholipase D generating anandamide and its congeners. J Biol Chem 279: 5298-5305. Onaivi ES, Chakrabarti A, Chaudhuri G. 1996. Cannabinoid receptor genes. Prog Neurobiol 48: 275-305. Onaivi ES, Leonard CM, Ishiguro H, Zhang PW, Lin Z, et al. 2002. Endocannabinoids and cannabinoid receptor genetics. Prog Neurobiol 66: 307-344.

Onaivi ES, Stubbs A, Chakrabarti L, Chittenden DP, Hurst BE, et al. 1998. Murine Cannabinoid receptor genetics. FASEB J 124: A194. Ortiz S, Oliva JM, Perez S, Palomo T, Manzanares J. 2004a. Chronic ethanol consumption regulates cannabinoid CB1 receptor gene expression in selected regions of rat brain. Alcohol 39: 88-92. Ortiz S, Oliva JM, Perez‐Rial S, Palomo T, Manzanares J. 2004b. Differences in basal cannabinoid CB1 receptor function in selective brain areas and vulnerability to voluntary alcohol consumption in Fawn Hooded and Wistar rats. Alcohol 39: 297-302. Palmer SL, Khanolkar AD, Makriyannis A. 2000. Natural and synthetic endocannabinoids and their structure ‐activity relationships. Curr Pharm Des 6: 1381-1397. Pan X, Ikeda SR, Lewis DL. 1996. Rat brain cannabinoid receptor modulates N‐type Ca2+ channels in a neuronal expression system. Mol Pharmacol 49: 707-714. Pan X, Ikeda SR, Lewis DL. 1998. SR 141716A acts as an inverse agonist to increase neuronal voltage‐dependent Ca2+ currents by reversal of tonic CB1 cannabinoid receptor activity. Mol Pharmacol 54: 1064-1072. Paria BC, Dey SK. 2000. Ligand–receptor signaling with endocannabinoids in preimplatation embryo development and implantation. Chem Phys Lipids 108: 211-220. Patricelli MP, Lovato MA, Cravatt BF. 1999. Chemical and mutagenic investigations of fatty acid amide hydrolase: Evidence for a family of serine hydrolases with distinct catalytic properties. Biochemistry 38: 9804-9812. Pertwee RG. 1997. Pharmacology of cannabinoid CB1 and CB2 receptors. Pharmacol Ther 74: 129-180. Pettit DA, Harrison MP, Olson JM, Spencer RF, Cabral GA. 1998. Immunohistochemical localization of the neural cannabinoid receptor in rat brain. J Neurosci Res 51: 391-402. Pfister‐Genskow M, Weesner GD, Hayes H, Eggen A, Bishop MD. 1997. Physical and genetic localization of the bovine cannabinoid receptor (CNR1) gene to bovine hromosome 9. Mamm Genome 8: 301-302. Pinto JC, Potie F, Rice KC, Boring D, Johnson MR, et al. 1994. Cannabinoid receptor binding and agonist activity of amides and esters of arachidonic acid. Mol Pharmacol 46: 516-522. Piomelli D. 2003. The molecular logic of endocannabinoid signalling. Nat Rev Neurosci 4: 873-884. Piomelli D, Beltramo M, Glasnapp S, Lin SY, Goutopoulos A, et al. 1999. Structural determinants for recognition and translocation by the anandamide transporter. Proc Natl Acad Sci USA 96: 5802-5807. Pitler TA, Alger BE. 1990. Activation of the pharmacologically defined M3 muscarinic receptor depolarizes hippocampal pyramidal cells. Brain Res 534: 257-262.

The endocannabinoid system Pitler TA, Alger BE. 1992. Postsynaptic spike firing reduces synaptic GABAA responses in hippocampal pyramidal cells. J Neurosci 12: 4122-4132. Pitler TA, Alger BE. 1994. Depolarization‐induced suppression of GABAergic inhibition in rat hippocampal pyramidal cells: G protein involvement in a presynaptic mechanism. Neuron 13: 1447-1455. Poncelet M, Maruani J, Calassi R, Soubrie P. 2003. Overeating, alcohol and sucrose consumption decrease in CB1 receptor deleted mice. Neurosci Lett 343: 216-218. Porter AC, Felder CC. 2001. The endocannabinoid nervous system: Unique opportunities for therapeutic intervention. Pharmacol Ther 90: 45-60. Porter AC, Sauer JM, Knierman MD, Becker GW, Berna MJ, et al. 2002. Characterization of a novel endocannabinoid, virodhamine, with antagonist activity at the CB1 receptor. J Pharmacol Exp Ther 301: 1020-1024. Prescott SM, Majerus PW. 1983. Characterization of 1,2‐diacylglycerol hydrolysis in human platelets. Demonstration of an arachidonoyl‐monoacylglycerol intermediate. J Biol Chem 258: 764-769. Racz I, Bilkei‐Gorzo A, Toth ZE, Michel K, Palkovits M, et al. 2003. A critical role for the cannabinoid CB1 receptors in alcohol dependence and stress‐stimulated ethanol drinking. J Neurosci 23: 2453-2458. Rakhshan F, Day TA, Blakely RD, Barker EL. 2000. Carrier‐ mediated uptake of the endogenous cannabinoid anandamide in RBL‐2H3 cells. J Pharmacol Exp Ther 292: 960-967. Ravinet Trillou C, Delgorge C, Menet C, Arnone M, Soubrie P. 2004. CB1 cannabinoid receptor knockout in mice leads to leanness, resistance to diet‐induced obesity and enhanced leptin sensitivity. Int J Obes Relat Metab Disord 28: 640-648. Ravinet Trillou C, Arnone M, Delgorge C, Gonalons N, Keane P, et al. 2003. Anti‐obesity effect of SR141716, a CB1 receptor antagonist, in diet‐induced obese mice. Am J Physiol Regul Integr Comp Physiol 284: R345-353. Reibaud M, Obinu MC, Ledent C, Parmentier M, Bohme GA, et al. 1999. Enhancement of memory in cannabinoid CB1 receptor knock‐out mice. Eur J Pharmacol 379: R1-2. Rhee MH, Bayewitch M, Avidor‐Reiss T, Levy R, Vogel Z. 1998. Cannabinoid receptor activation differentially regulates the various adenylyl cyclase isozymes. J Neurochem 71: 1525-1534. Rimondini R, Arlinde C, Sommer W, Heilig M. 2002. Long‐ lasting increase in voluntary ethanol consumption and transcriptional regulation in the rat brain after intermittent exposure to alcohol. FASEB J 16: 27-35. Robbe D, Kopf M, Remaury A, Bockaert J, Manzoni OJ. 2002. Endogenous cannabinoids mediate long‐term synaptic

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depression in the nucleus accumbens. Proc Natl Acad Sci USA 99: 8384-8388. Robbins TW, Everitt BJ. 1996. Neurobehavioural mechanisms of reward and motivation. Curr Opin Neurobiol 6: 228-236. Rodriguez de Fonseca F, Roberts AJ, Bilbao A, Koob GF, Navarro M. 1999. Cannabinoid receptor antagonist SR 141716A decreases operant ethanol self administration in rats exposed to ethanol‐vapor chambers. Acta Pharmacol Sin 20: 1109-1114. Rosenberg CM. 1979. Alcoholics anonymous as treatment and as ideology; comments on the article by R. E. Tournier. J Stud Alcohol 40: 330-333. Rosenberg CM, Gerrein JR, Schnell C. 1978. Cannabis in the treatment of alcoholism. J Stud Alcohol 39: 1955-1958. Rowland RS. 2002. Using X‐ray crystallography in drug discovery. Curr Opin Drug Discov Devel 5: 613-619. Rueda D, Galve‐Roperh I, Haro A, Guzman M. 2000. The CB(1) cannabinoid receptor is coupled to the activation of c‐Jun N‐terminal kinase. Mol Pharmacol 58: 814-820. Ryan WJ, Banner WK, Wiley JL, Martin BR, Razdan RK. 1997. Potent anandamide analogs: The effect of changing the length and branching of the end pentyl chain. J Med Chem 40: 3617-3625. Ryberg E, Vu HK, Larsson N, Groblewski T, Hjorth S, et al. 2005. Identification and characterisation of a novel splice variant of the human CB1 receptor. FEBS Lett 579: 259-264. Salzet M, Breton C, Bisogno T, Di Marzo V. 2000. Comparative biology of the endocannabinoid system. Possible role in the immune response. Eur J Biochem 267: 4917-4927. Sanchez C, Galve‐Roperh I, Rueda D, Guzman M. 1998. Involvement of sphingomyelin hydrolysis and the mitogen‐activated protein kinase cascade in the delta9‐tetrahydrocannabinol‐induced stimulation of glucose metabolism in primary astrocytes. Mol Pharmacol 54: 834-843. Sanchez C, Rueda D, Segui B, Galve‐Roperh I, Levade T, et al. 2001. The CB(1) cannabinoid receptor of astrocytes is coupled to sphingomyelin hydrolysis through the adaptor protein fan. Mol Pharmacol 59: 955-959. Schabitz WR, Giuffrida A, Berger C, Aschoff A, Schwaninger M, et al. 2002. Release of fatty acid amides in a patient with hemispheric stroke: A microdialysis study. Stroke 33: 2112-2114. Schmid PC, Reddy PV, Natarajan V, Schmid HH. 1983. Metabolism of N‐acylethanolamine phospholipids by a mammalian phosphodiesterase of the phospholipase D type. J Biol Chem 258: 9302-9306. Schultes RE. 1967. Man and marijuana. Nat Hist 82: 59-63, 80, 82. Schweitzer P. 2000. Cannabinoids decrease the K(+) M‐current in hippocampal CA1 neurons. J Neurosci 20: 51-58.

381

382

14

The endocannabinoid system

Self DW. 1999. Anandamide: A candidate neurotransmitter heads for the big leagues. Nat Neurosci 2: 303-304. Seltzman HH, Fleming DN, Thomas BF, Gilliam AF, McCallion DS, et al. 1997. Synthesis and pharmacological comparison of dimethylheptyl and pentyl analogs of anandamide. J Med Chem 40: 3626-3634. Serra S, Brunetti G, Pani M, Vacca G, Carai MA, et al. 2002. Blockade by the cannabinoid CB(1) receptor antagonist, SR 141716, of alcohol deprivation effect in alcohol‐preferring rats. Eur J Pharmacol 443: 95-97. Shen M, Thayer SA. 1998. The cannabinoid agonist Win55, 212‐2 inhibits calcium channels by receptor‐mediated and direct pathways in cultured rat hippocampal neurons. Brain Res 783: 77-84. Sim‐Selley LJ, Martin BR. 2002. Effect of chronic administration of R‐(+)‐[2,3‐Dihydro‐5‐methyl‐3‐[(morpholinyl) methyl]pyrrolo[1,2,3‐de]‐1,4‐benzoxazinyl]‐(1‐naphthalenyl)methanone mesylate (WIN55, 212‐2) or delta (9)‐tetrahydrocannabinol on cannabinoid receptor adaptation in mice. J Pharmacol Exp Ther 303: 36-44. Sipe JC, Chiang K, Gerber AL, Beutler E, Cravatt BF. 2002. A missense mutation in human fatty acid amide hydrolase associated with problem drug use. Proc Natl Acad Sci USA 99: 8394-8399. Soderstrom K, Johnson F. 2000. CB1 cannabinoid receptor expression in brain regions associated with zebra finch song control. Brain Res 857: 151-157. Stanton PK, Sarvey JM. 1985. Depletion of norepinephrine, but not serotonin, reduces long‐term potentiation in the dentate gyrus of rat hippocampal slices. J Neurosci 5: 2169-2176. Stefano GB, Salzet B, Salzet M. 1997. Identification and characterization of the leech CNS cannabinoid receptor: Coupling to nitric oxide release. Brain Res 753: 219-224. Steiner H, Bonner TI, Zimmer AM, Kitai ST, Zimmer A. 1999. Altered gene expression in striatal projection neurons in CB1 cannabinoid receptor knockout mice. Proc Natl Acad Sci USA 96: 5786-5790. Stella N, Piomelli D. 2001. Receptor‐dependent formation of endogenous cannabinoids in cortical neurons. Eur J Pharmacol 425: 189-196. Stella N, Schweitzer P, Piomelli D. 1997. A second endogenous cannabinoid that modulates long‐term potentiation. Nature 388: 773-778. Stubbs L, Chittenden L, Chakrabarti A, Onaivi E. 1996. The gene encoding the central cannabinoid receptor is located in proximal mouse Chromosome 4. Mamm Genome 7: 165-166. Sugiura T, Kobayashi Y, Oka S, Waku K. 2002. Biosynthesis and degradation of anandamide and 2‐arachidonoylglycerol and their possible physiological significance. Prostaglandins Leukot Essent Fatty Acids 66: 173-192.

Sugiura T, Yoshinaga N, Kondo S, Waku K, Ishima Y. 2000. Generation of 2‐arachidonoylglycerol, and endogenous cannabinoid receptor ligand, in picrotoxinin‐ administered rat brain. Biochem Biophys Res Commun 271: 654-658. Sugiura T, Kodaka T, Kondo S, Nakane S, Kondo H, et al. 1997. Is the cannabinoid CB1 receptor a 2‐arachidonoylglycerol receptor? Structural requirements for triggering a Ca2þ transient in NG108–15 cells. J Biochem (Tokyo) 122: 890-895. Sugiura T, Kodaka T, Nakane S, Kishimoto S, Kondo S, et al. 1998. Detection of an endogenous cannabimimetic molecule, 2‐arachidonoylglycerol, and cannabinoid CB1 receptor mRNA in human vascular cells: Is 2‐arachidonoylglycerol a possible vasomodulator? Biochem Biophys Res Commun 243: 838-843. Sugiura T, Kodaka T, Nakane S, Miyashita T, Kondo S, et al. 1999. Evidence that the cannabinoid CB1 receptor is a 2‐arachidonylglycerol receptor. J Biol Chem 274: 2794-2801. Sugiura T, Kondo S, Sukagawa A, Nakane S, Shinoda A, et al. 1995. 2‐Arachidonoylglycerol: A possible endogenous cannabinoid receptor ligand in brain. Biochem Biophys Res Commun 215: 89-97. Sugiura T, Kondo S, Sukagawa A, Tonegawa T, Nakane S, et al. 1996a. N‐arachidonoylethanolamine (anandamide), an endogenous cannabinoid receptor ligand, and related lipid molecules in the nervous tissues. J Lipid Mediat Cell Signal 14: 51-56. Sugiura T, Kondo S, Sukagawa A, Tonegawa T, Nakane S, et al. 1996b. Enzymatic synthesis of anandamide, an endogenous cannabinoid receptor ligand, through N‐acylphosphatidylethanolamine pathway in testis: Involvement of Ca(2+)‐ dependent transacylase and phosphodiesterase activities. Biochem Biophys Res Commun 218: 113-117. Sugiura T, Kodaka T, Kondo S, Tonegawa T, Nakane S, et al. 1996c. 2‐Arachidonoylglycerol, a putative endogenous cannabinoid receptor ligand, induces rapid, transient elevation of intracellular free Ca2+ in neuroblastoma x glioma hybrid NG108–15 cells. Biochem Biophys Res Commun 229: 58-64. Sullivan JM. 2000. Cellular and molecular mechanisms underlying learning and memory impairments produced by cannabinoids. Learn Mem 7: 132-139. Szabo B, Muller T, Koch H. 1999. Effects of cannabinoids on dopamine release in the corpus striatum and the nucleus accumbens in vitro. J Neurochem 73: 1084-1089. Szabo B, Wallmichrath I, Mathonia P, Pfreundtner C. 2000. Cannabinoids inhibit excitatory neurotransmission in the substantia nigra pars reticulata. Neuroscience 97: 89-97. Szabo B, Dorner L, Pfreundtner C, Norenberg W, Starke K. 1998. Inhibition of GABAergic inhibitory postsynaptic

The endocannabinoid system currents by cannabinoids in rat corpus striatum. Neuroscience 85: 395-403. Takahashi KA, Linden DJ. 2000. Cannabinoid receptor modulation of synapses received by cerebellar Purkinje cells. J Neurophysiol 83: 1167-1180. Tanda G, Pontieri FE, Di Chiara G. 1997. Cannabinoid and heroin activation of mesolimbic dopamine transmission by a common mul opioid receptor mechanism. Science 276: 2048-2050. Tart CT. 1970. Marijuana intoxication common experiences. Nature 226: 701-704. Thomas BF, Adams IB, Mascarella SW, Martin BR, Razdan RK. 1996. Structure‐activity analysis of anandamide analogs: Relationship to a cannabinoid pharmacophore. J Med Chem 39: 471-479. Tsou K, Brown S, Sanudo‐Pena MC, Mackie K, Walker JM. 1998a. Immunohistochemical distribution of cannabinoid CB1 receptors in the rat central nervous system. Neuroscience 83: 393-411. Tsou K, Nogueron MI, Muthian S, Sanudo‐Pena MC, Hillard CJ, et al. 1998b. Fatty acid amide hydrolase is located preferentially in large neurons in the rat central nervous system as revealed by immunohistochemistry. Neurosci Lett 254: 137-140. Twitchell W, Brown S, Mackie K. 1997. Cannabinoids inhibit N‐ and P/Q‐type calcium channels in cultured rat hippocampal neurons. J Neurophysiol 78: 43-50. Ueda N, Kurahashi Y, Yamamoto S, Tokunaga T. 1995. Partial purification and characterization of the porcine brain enzyme hydrolyzing and synthesizing anandamide. J Biol Chem 270: 23823-23827. Ueda N, Puffenbarger RA, Yamamoto S, Deutsch DG. 2000. The fatty acid amide hydrolase (FAAH). Chem Phys Lipids 108: 107-121. Valjent E, Pages C, Rogard M, Besson MJ, Maldonado R, et al. 2001. Delta 9‐tetrahydrocannabinol‐induced MAPK/ERK and Elk‐1 activation in vivo depends on dopaminergic transmission. Eur J Neurosci 14: 342-352. Valk PJ, Hol S, Vankan Y, Ihle JN, Askew D, et al. 1997. The genes encoding the peripheral cannabinoid receptor and alpha‐L‐fucosidase are located near a newly identified common virus integration site, evill. J Virol 71: 6796-6804. Valverde O, Noble F, Beslot F, Dauge V, Fournie‐Zaluski MC, et al. 2001. Delta 9‐tetrahydrocannabinol releases and facilitates the effects of endogenous enkephalins: Reduction in morphine withdrawal syndrome without change in rewarding effect. Eur J Neurosci 13: 1816-1824. Varma N, Carlson GC, Ledent C, Alger BE. 2001. Metabotropic glutamate receptors drive the endocannabinoid system in hippocampus. J Neurosci 21: RC188. Vaughan CW, McGregor IS, Christie MJ. 1999. Cannabinoid receptor activation inhibits GABAergic neurotransmission

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in rostral ventromedial medulla neurons in vitro. Br J Pharmacol 127: 935-940. Vaughan CW, Connor M, Bagley EE, Christie MJ. 2000. Actions of cannabinoids on membrane properties and synaptic transmission in rat periaqueductal gray neurons in vitro. Mol Pharmacol 57: 288-295. Vincent P, Marty A. 1993. Neighboring cerebellar Purkinje cells communicate via retrograde inhibition of common presynaptic interneurons. Neuron 11: 885-893. Vincent P, Marty A. 1996. Fluctuations of inhibitory postsynaptic currents in Purkinje cells from rat cerebellar slices. J Physiol 494 (Pt 1): 183-199. Vincent P, Armstrong CM, Marty A. 1992. Inhibitory synaptic currents in rat cerebellar Purkinje cells: Modulation by postsynaptic depolarization. J Physiol 456: 453-471. Wagner JA, Varga K, Jarai Z, Kunos G. 1999. Mesenteric vasodilation mediated by endothelial anandamide receptors. Hypertension 33: 429-434. Walker JM, Huang SM. 2002. Endocannabinoids in pain modulation. Prostaglandins Leukot Essent Fatty Acids 66: 235-242. Wallmichrath I, Szabo B. 2002. Cannabinoids inhibit striatonigral GABAergic neurotransmission in the mouse. Neuroscience 113: 671-682. Wang L, Liu J, Harvey‐white J, Zimmer A, Kunos G. 2003. Endocannabinoid signaling via CB1 receptors is involved in ethanol preference and its age‐dependent decline in mice. Proc Natl Acad Sci USA 100: 1393-1398. Wartmann M, Campbell D, Subramanian A, Burstein SH, Davis RJ. 1995. The MAP kinase signal transduction pathway is activated by the endogenous cannabinoid anandamide. FEBS Lett 2‐3: 133-136. Willoughby KA, Moore SF, Martin BR, Ellis EF. 1997. The biodisposition and metabolism of anandamide in mice. J Pharmacol Exp Ther 282: 243-247. Wilson RI, Nicoll RA. 2001. Endogenous cannabinoids mediate retrograde signalling at hippocampal synapses. Nature 410: 588-592. Wilson RI, Nicoll RA. 2002. Endocannabinoid Signaling in the Brain. Science 296: 678-682. Wilson RI, Kunos G, Nicoll RA. 2001. Presynaptic specificity of endocannabinoid signaling in the hippocampus. Neuron 31: 453-462. Wise RA. 1996. Addictive drugs and brain stimulation reward. Annu Rev Neurosci 19: 319-340. Wise RA, Bozarth MA. 1987. A psychomotor stimulant theory of addiction. Psychol Rev 94: 469-492. Yamaguchi F, Macrae AD, Brenner S. 1996. Molecular cloning of two cannabinoid type 1‐like receptor genes from the puffer fish Fugu rubripes. Genomics 35: 603-605. Yang HY, Karoum F, Felder C, Badger H, Wang TC, et al. 1999. GC/MS analysis of anandamide and quantification of

383

384

14

The endocannabinoid system

N‐arachidonoylphosphatidylethanolamides in various brain regions, spinal cord, testis, and spleen of the rat. J Neurochem 72: 1959-1968. Yoshida T, Hashimoto K, Zimmer A, Maejima T, Araishi K, et al. 2002. The cannabinoid CB1 receptor mediates retrograde signals for depolarization‐induced suppression of inhibition in cerebellar Purkinje Cells. J Neurosci 22: 1690-1697. Zhang PW, Ishiguro H, Ohtsuki T, Hess J, Carillo F, et al. 2004. Human cannabinoid receptor 1:50 exons, candidate

regulatory regions, polymorphisms, haplotypes and association with polysubstance abuse. Mol Psychiatry 9: 916-931. Zilberter Y. 2000. Dendritic release of glutamate suppresses synaptic inhibition of pyramidal neurons in rat neocortex. J Physiol 528 (Pt 3): 489-496. Zimmer A, Zimmer AM, Hohmann AG, Herkenham M, Bonner TI. 1999. Increased mortality, hypoactivity, and hypoalgesia in cannabinoid CB1 receptor knockout mice. Proc Natl Acad Sci USA 96: 5780-5785.

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E Prostanoid Receptors in Brain Physiology and Disease

C. D. Keene . P. J. Cimino . R. M. Breyer . K. S. Montine . T. J. Montine

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Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 386

2 2.1 2.2 2.3 2.4

Prostaglandin Pathway and E Prostanoid (EP) Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 386 Eicosanoids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 386 Prostaglandin Pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 386 EP Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 387 Expression of EP Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 387

3 3.1 3.2 3.2.1 3.2.2 3.2.3 3.2.4

Physiologic and Pathophysiologic Roles for EP Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 388 Periphery . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 388 Central Nervous System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 390 Excitotoxicity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 391 Aggregated Ab42 Neurotoxicity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 392 Innate Immune Response . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 392 Microglia Phagocytosis of Neurotoxic Peptides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 393

4 4.1 4.2 4.2.1 4.2.2 4.2.3 4.2.4

Human Brain Diseases and Their Models . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 394 Human Neurologic Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 394 Disease Models . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 394 Ischemia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 394 Alzheimer’s Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 395 Parkinson’s Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 395 Amyotrophic Lateral Sclerosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 396

5

Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 396

#

2008 Springer ScienceþBusiness Media, LLC.

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Abstract: Here we will review the prostaglandin (PG) pathway with a focus on emerging data that highlight a central role for PGE2 in ischemic injury and several neurodegenerative diseases. PGE2 activates a family of four G protein‐coupled receptors, EP1 to EP4, which are expressed throughout the central nervous system on different cell types where they play central roles in physiologic and pathophysiologic responses. Of these, the data most clearly point to EP1 activation as a mediator of neurotoxicity in models of ischemic injury and EP2 activation as a mediator of activated microglial paracrine damage to neurons and a suppressor of microglia phagocytosis of aggregated neurotoxic peptides. Preclinical animal models of some human neurologic diseases support the potential efficacy of targeting specific EP receptor subtypes in Alzheimer’s disease, Parkinson’s disease, amyotrophic lateral sclerosis, and ischemic stroke. Clinical trials targeting any EP receptor in neurologic disease have not yet been reported. List of Abbreviations: AA, arachidonic acid; AD, Alzheimer’s disease; ALS, amyotrophic lateral sclerosis; APP, amyloid precursor protein; CNS, central nervous system; COXs, cyclooxygenases; CRTH2, chemoattractant receptor homologous molecule expressed on Th2 cells; CSF, cerebrospinal fluid; DHA, docosahexaenoic acid; FAD, Familial Alzheimer’s disease; ICAM, intercellular adhesion molecule; ICV, intracerebroventricular; iNOS, inducible nitric oxide synthase; IL‐6, interleukin‐6; LGs, Levuglandins; LOXs, lipoxygenases; LPS, lipopolysaccharide; MCP‐1, monocyte chemotactic protein‐1; MPTP, 1‐methyl‐ 4‐phenyl‐1,2,3,6‐tetrahydropyridine; NMDA, N‐methyl‐D‐aspartic acid; NSAIDs, nonsteroidal anti‐ inflammatory drugs; PD, Parkinson’s disease; PG, prostaglandin; PL, phospholipase; PS, presenilin; ROS, reactive oxygen species; SOD1, superoxide dismutase; TNF‐a, tumor necrosis factor‐a; Tx, thromboxane

1

Introduction

Prostaglandins (PGs) are potent autocrine and paracrine factors that contribute significantly to homeostasis as well as disease pathogenesis in virtually every organ. Here we will review the PG pathway and then focus on emerging data that highlight a central role for one PG in neurologic diseases.

2

Prostaglandin Pathway and E Prostanoid (EP) Receptors

2.1 Eicosanoids Eicosanoids are oxygenated products of arachidonic acid (AA). The ester linkage that binds AA to the glycerol backbone of phospholipids is hydrolyzed by phospholipase (PL) A2 in response to a wide array of physiologic and pathophysiologic stimuli (Kudo and Murakami, 2002). There are two predominant groups of enzymes that catalyze oxygenation of the hydrolyzed AA to yield precursors of potent bioactive products. The first group is the cyclooxygenases (COXs), which catalyze the formation of prostaglandin (PG) H2 from free AA. The second is the lipoxygenases (LOXs), which catalyze the formation of hydroperoxides from AA and likely also docosahexaenoic acid (DHA) (Campbell and Halushka, 2001). While cytochrome P450s also catalyze oxygenation of AA, the bioactivity of these compounds is less clear.

2.2 Prostaglandin Pathway Isozymes of COX catalyze oxygenation of hydrolyzed AA in what is the committed step in PG synthesis (Kaufmann et al., 1997). A bis‐oxygenase reaction in which AA plus two O2 are converted to the bicycloendoperoxide intermediate PGG2 is followed by a peroxidase step to generate PGH2 with the release of an oxidizing radical (Siedlik and Marnett, 1984). PGH2 exerts biological activity through three different mechanisms. First, PGH2 may be further metabolized by a number of cell‐specific synthases or isomerases to PGD2, PGE2, PGF2a, PGI2, and thromboxane (Tx) A2, all of which exert potent autocrine and paracrine activities through the stimulation of eight specific cell surface rhodopsin‐like transmembrane spanning

E prostanoid receptors in brain physiology and disease

15

receptors (GPCRs) designated DP, EP1–4, FP, IP, and TP, respectively (Hata and Breyer, 2004). Recently, a ninth prostaglandin receptor was identified—the chemoattractant receptor homologous molecule expressed on Th2 cells (CRTH2)—that binds PGD2 (Hirai et al., 2001). It is through the coordinated cell‐specific expression of these synthases or isomerases and receptors that products of COX achieve such a wide variety of actions in different cells and tissue. Second, PGH2 itself is an agonist for TP. Finally, PGH2 may rearrange to form levuglandins (LGs), highly chemically reactive g‐ketoaldehydes, which form irreversible adducts with e‐amino groups of protein lysyl residues leading to protein–protein crosslinks (Difranco et al., 1994); indeed, LGs significantly accelerate oligomerization of Ab peptides in vitro (Boutaud et al., 2002).

2.3 EP Receptors Although similar in some aspects, PGE2 is distinct from other eicosanoid products of COX because of the existence of widely expressed multiple receptor subtypes, EP1, EP2, EP3, and EP4, linked to functionally antagonistic second messenger systems; because of this PGE2 has versatile and often opposing actions in tissues and cells [reviewed in (Breyer et al., 2001)]. The precise roles of EP receptors in physiologic and pathologic settings are determined by multiple factors such as ligand affinity, receptor expression profile, differential coupling to signal transduction pathways, and the cellular context in which the receptor is expressed. EP3 is unlike the other EP receptors in that it has three different splice variants, EP3a, EP3b, and EP3g; EP3b is unique as it does not desensitize and thus displays persistent signaling when exposed to ligand (Negishi et al., 1993; Breyer et al., 2001). Some characteristics of EP receptors and commonly used agents are presented in > Table 15‐1.

. Table 15‐1 Characteristics of EP receptor subtypes

EP1

Ki PGE2 (nM) human (mouse) 9.1 (20)

G‐protein coupling unknown

EP2

4.9 (12)

Gs

EP3

0.3 (0.9)

Gi Gq Gs

EP4

0.8 (1.9)

Gs

Signaling ↑ Ca2þ

↑ cAMP, EGFR transactivation b‐catenin ↓ cAMP ↑ IP3/DAG ↑ cAMP ↑ cAMP, PI3K, ERK1/2, b‐catenin

Relatively selective agonists ONO‐KI‐004, iloprost, 17‐phenyl‐trinor PGE2, sulprostone Butaprost, 11‐deoxy PGE1, AH13205, ONO‐AEI‐259 Sulprostone, MB28767, misoprostol, SC46275, ONO‐AE‐249 PGE1‐OH, misoprostol, ONO‐AEI‐329

Relatively selective antagonists SC51322, SC51089, ONO‐8713 NONE

ONO‐AE3‐ 240, L‐826266

AH23848B ONO‐ AE3‐208

2.4 Expression of EP Receptors Restricted expression of EP receptors in different organs and even different cells within an organ is one of the principal means by which PGE2 signaling attains such varied outcomes [reviewed in (Breyer et al., 2001)]. Pertinent to our review of EP receptors in the brain, there is some controversy over the expression of EP receptor subtypes in the central nervous system (CNS), which appears to derive from technical issues related to the specificity of commercially available antibodies and the purity of some primary cultures used

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for PCR. Antibodies currently available to EP2 and EP3 receptor subtypes are not specific and highlight many bands from brain homogenates probed by Western blot (unpublished data). Thus, we are skeptical not only of Western blot results using these antibodies but especially conclusions drawn from immunohistochemical studies. Indeed, we have been able to reproduce the findings of others’ immunohistochemical analysis of brain using antibodies to EP2; however, we obtain the same immunoreactivity in EP2
/
mice, a control not run by others. Here we will broadly summarize the studies presented in > Table 15‐2. All of the EP receptors are expressed in rodent brain, where there are regional and cell‐specific differences in expression and activity. In situ probing c‐fos expression experiments in rats have shown that EP1, EP2, and EP3 are expressed by neurons in multiple regions of brain (Sugimoto et al., 1994; Bhattacharya et al., 1998; Ek et al., 2000; Nakamura et al., 2000; Oka et al., 2000; Kawano et al., 2006). Rodent neuronal EP4 expression is more restricted to some hypothalamic nuclei (Zhang and Rivest, 1999). Expression of all EP receptor subtypes has been reported in primary culture of either microglia or astrocytes; however, not all results have been reproducible. When interpreting these experiments, it is important to keep in mind that culturing of glia from brain results in some level of activation that is not present in vivo. EP2 is functionally expressed on microglia in culture and on activated microglia in vivo. EP1 and (inducible) EP3 may be expressed on microglia, although we are unaware of any data showing activity; indeed, two groups have not observed modification of microglial response to LPS in EP1
/
microglia (Milatovic et al., 2005; Kawano et al., 2006). EP2, (inducible) EP3, and EP4 appear to be expressed by astrocytes.

3

Physiologic and Pathophysiologic Roles for EP Receptors

3.1 Periphery PGE2 is a major product of the PG pathway in a number of physiologic and pathologic settings where it plays a critical role in maintaining the integrity of gastric mucosa (Woo et al., 1986; Warner et al., 1999), preserving renal blood flow and glomerular filtration rate (for a review, see Breyer and Breyer, 2001), and maintaining blood pressure (Kennedy et al., 1999). PGE2 has been observed to have multiple and apparently opposing functional effects. For example, PGE2 elicits both smooth muscle relaxation and constriction in the respiratory tree and in vascular smooth muscle (Gardiner, 1986; Walch et al., 2001; Davis et al., 2004). PGE2 activation of EP receptors is critical to several aspects of the innate and adaptive immune response in the periphery. EP1 plays a role in inflammatory hyperalgesia (Stock et al., 2001; Nakayama et al., 2002). EP2 contributes to inhibition of T‐cell proliferation (Nataraj et al., 2001), IgE class switching by B‐cells (Fedyk and Phipps, 1996), and inhibition of dendritic cell function in cancer‐associated immunodeficiency (Yang et al., 2003). EP3, like EP1, is involved in inflammatory hyperalgesia (Minami et al., 2001) as well as enhancement of antigen‐stimulated mast cell degranulation (Nguyen et al., 2002). Finally, EP4 signaling contributes to enhanced migration, maturation, and T‐cell stimulatory capacity of Langerhans cells (Kabashima et al., 2003). EP2 has been studied extensively with regard to innate immunity in the periphery. Expression of peripheral macrophage EP2 is upregulated on antigen‐presenting cells in models of innate immunity (Hubbard et al., 2001; Harizi et al., 2003), and EP2 regulates expression of inflammatory mediators, including tumor necrosis factor‐ (TNF‐) a (Fennekohl et al., 2002; Vassiliou et al., 2003; Akaogi et al., 2004), interleukin‐ (IL‐) 6 (Akaogi et al., 2004; Treffkorn et al., 2004), monocyte chemotactic protein‐ (MCP‐) 1 (Largo et al., 2004), intercellular adhesion molecule (ICAM) 1 (Noguchi et al., 2001), and inducible nitric oxide synthase (iNOS) (Minghetti et al., 1997). Separate studies point to additional roles of EP2 receptor in modulating macrophage migration (Baratelli et al., 2004) and inhibiting phagocytosis of bacterial components by lung alveolar macrophages (Aronoff et al., 2004). These studies are consistent with others that demonstrate that PGE2 inhibits phagocytosis by macrophages by a process that is dependent on increased cAMP levels (Hutchison and Myers, 1987; Canning et al., 1991; Borda et al., 1998; Aronoff et al., 2004). Conversely, nonsteroidal anti‐inflammatory drugs (NSAIDs) potentiate phagocytosis by macrophages (Bjornson et al., 1988; Gilmour et al., 1993; Gurer et al., 2002).

E prostanoid receptors in brain physiology and disease

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. Table 15‐2 Anatomic, cellular, and subcellular localization of EP receptor subtypes Species Technique Anatomic localization Rat PCR, WB, and IHC

Receptor

Results

References

EP1

Most abundantly expressed in parietal cortex and cerebellum where IHC localized to Purkinje cells. Widely expressed on neurons in cerebral cortex Predominantly expressed in hypothalamus Brain. Bed nucleus of the stria terminalis, lateral septum, subfornical organ, ventromedial hypothalamic nucleus, central nucleus of the amygdala, locus coeruleus (LC), and the area postrema Brain. Preoptic area, magnocellular paraventricular nucleus, supraoptic nucleus, parabrachial (PB) nucleus, LC, nucleus of the solitary tract (NTS), and ventrolateral medulla (VLM) Widespread on neurons throughout brain PB. Preprodynorphin‐expressing neurons in the dorsal and central lateral subnuclei; preproenkephalin‐, calcitonin gene‐related peptide‐, and preprotachykinin‐expressing neurons in the external lateral subnucleus, and some enkephalinergic neurons in Kolliker‐Fuse nucleus PB. Neurons in central cholecystokininergic population of superior lateral subnucleus Neuraxis. Olfactory system, iso‐ and hippocampal cortices, subcortical telencephalic structures in the septal region and amygdale, Thalamic groups (midline, intralaminar, and anterior), preoptic nuclei, brainstem periaqueductal gray (PAG), LC, PB, raphe, NTS, and VLM Neuraxis. Especially IR were thalamic groups (anterior, intralaminar, and midline) preoptic nucleus medial mammillary nucleus, superior colliculus, PAG, LC, PB, raphe, NTS, laminae I and II of the medullary and spinal dorsal horns, and sensory ganglia

Candelario‐Jalil et al. (2005)

Mouse

IHC

EP1

Mouse

ISH

EP1

Rat

ISH

EP2

EP4

Mouse

ISH

EP3

Rat

ISH

EP3

EP4

Rat

ISH

EP3

Rat

IHC

EP3

Kawano et al. (2006) Batshake et al. (1995) Zhang and Rivest (1999)

Sugimoto et al. (1994) Engblom et al. (2004)

Ek et al. (2000)

Nakamura et al. (2000)

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. Table 15‐2 (continued) Species Rat

Technique IHC

Receptor EP3

Cellular and subcellular localization Rat PCR EP1, EP2, microglia EP3, and EP4 Rat WB and EP2 and IHC EP4

Rat

WB, ICC, RNAi

EP2 and EP3

Rat

IHC

Rat microglia or astrocytes Rat striatum

PCR

EP2 and EP3 EP3

IHC

EP3

Pig or rat brain

RB, IF, EM

EP3 and EP4

Human or rat astrocytes

PCR

EP2, EP3, and EP4

Results Brainstem serotonergic and catecholaminergic groups. Serotonergic: almost all in the medulla oblongata (B1–B4) with fewer in mesencephalic and pontine groups (B5–B9). Catecholaminergic: many of the noradrenergic A7 cells, LC densely in the neuropil and occasionally in neurons. No IR in noradrenergic A2 and A4, the adrenergic C2, and all the dopaminergic cell groups

References Nakamura et al. (2001)

Only EP1 and EP2 expressed

Caggiano and Kraig (1999)

Increased in reactive glial cells in CA1 and hilar regions of rats subjected to lethal ischemia without preconditioning. Most EP2 IR localized to microglia, whereas EP4 IR was found only on astrocytes Bidirectional activity‐dependent receptor trafficking at post‐synaptic membranes in visual cortex slices contributes to long‐term potentiation Ventral Spinal Cord. Motor neurons and astrocytes Both types of cells in culture expressed EP3

Choi et al. (2006)

Delayed appearance or EP3 IHC on microglia following excitotoxic lesion Present in nuclear membrane preparation and localized to nuclear envelope EP2 and EP4 expression in basal cultures. IL‐1b‐induced expression of EP3

Akaneya and Tsumoto (2006)

Bilak et al. (2004) Kitanaka et al. (1996)

Slawik et al. (2004) Bhattacharya (1998)

Waschbisch et al. (2006)

Abbreviations: EM, electron microscopy; ICC, immunocytochemistry; IF, immunofluorescence; IHC, immunohistochemistry; ISH, in situ hybridization; PCR, polymerase chain reaction; RNAi, RNA interference; RB, radioligand binding; WB, Western blot

3.2 Central Nervous System Research in several laboratories continues to clarify the role of PGE2 and specific EP receptors in CNS physiology. For example, PGE2 plays a central role in stimulated release of glutamate from astrocytes, a process proposed to be critical to neuron–glia communication at the synapse (Bezzi et al., 1998), whereas

E prostanoid receptors in brain physiology and disease

15

complex opposing roles for EP2 and EP3 receptors have been proposed in long‐term potentiation in primary visual cortex (Akaneya and Tsumoto, 2006). Here we will focus on the contribution of PGE2 signaling to mechanisms that mediate CNS injury (> Table 15‐3). These mechanisms include both direct damage to neurons from challenge with excitotoxins or aggregated Ab peptides and glial activities that, depending on the context of stressors, can produce paracrine damage to neurons through products of innate immunity or can promote neuronal survival by modulating phagocytosis of neurotoxic peptides.

3.2.1 Excitotoxicity Excitatory neurotransmission is mediated largely by synaptic glutamate activating one of several receptors that are named after their selective pharmacologic ligands; one is the N‐methyl‐D‐aspartic acid (NMDA) receptor, a ligand‐gated ion channel with high Ca2þ capacitance, whereas another ionotropic glutamate receptor is the kainate receptor (Planells‐Cases et al., 2006). COX‐2 neuronal expression is tightly regulated by excitatory synaptic activity via the NMDA receptor (Yamagata et al., 1993), and therefore many laboratories have explored the mechanistic links between PG signaling and excitatory neurotransmission (see earlier for examples). In the process known as excitotoxicity, protracted high levels of synaptic glutamate lead to excessive activation of NMDA and other glutamate receptors to culminate in relatively selective neuron damage and

. Table 15‐3 EP receptors in neuronal and glial mechanisms of neurologic disease

Neuron

Astrocyte

Microglia

EP1 activation Promotec (D: Mu C)

Mechanism/effect Excitotoxin‐induced neuron damage

NSAIDs Suppressa,b (D: Mu N, Rt H)

Ab‐induced neuron damage or death

Suppressg–j (D: Mu)

???

TNF‐a‐mediated glutamate release PGE2‐stimulated IL‐6 secretion LPS or Ab‐induced cytokine secretion

Suppressm (D: H and Rt A) ???

Not expressed No effectn (D: Rt A) ???

LPS or Ab‐induced paracrine neuron damage or death Phagocytosis of Ab or a‐synuclein

Suppressp (D: Mu M, C)

Suppress (D: Mu M, OH)

No effectt (D: Mu M)

No effectc,q (KO: Mu C; D: Mu OH) ???

EP2 activation Suppressd,e (D: Rt OH, OSC) No Effectk (KO: Mu N) Suppressl (D: Mu N) ??? No effectn (D: Rt A) Promotek,o (KO: Mu M, OH) Promotek,o,q–s (KO: Mu M, A, CC) Suppressk,u (KO: Mu M)

EP3 activation ???

EP4 activation Suppressf (D: Mu C)

???

Suppressl (D: Mu N)

???

???

No effectn (D: Rt A) ???

Presumedn (Rt A) Not expressed

???

Not expressed

???

Not expressed

Abbreviations: drug (D), gene knockout (KO); mouse (Mu), rat (Rt); primary neuron cultures (N), primary astrocyte cultures (A), primary microglia cultures (M), organotypic hippocampal cultures (OH), organotypic spinal cord cultures (OSC), cerebrum in vivo (C), hippocampus in vivo (H). aHewett et al. (2000); bPepicelli et al. (2005); cKawano et al. (2006); dLiu et al. (2005); eBilak et al. (2004); fAhmad et al. (2005); gLim et al. (2000); hLim et al. (2001); iJantzen et al. (2002); jYan et al. (2003); kShie et al. (2005a); lEcheverria et al. (2005); mBezzi et al. (2001); nFiebich et al. (2001); oShie et al. (2005b); pMilatovic et al. (2003); qMilatovic et al. (2005); rMontine et al. (2002); sMilatovic et al. (2004); tunpublished data; uJin et al. (2007); ???—not yet investigated

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ultimately death. Excitotoxicity has been proposed to contribute to many neurologic diseases including ischemic damage, traumatic brain injury, and virtually every type of neurodegenerative disease. Several groups have investigated the contribution of the PG pathway and specific EP receptors in models where CNS neurons are challenged with excitotoxin exposure (> Table 15‐3). Inhibition of COX activity, predominantly COX‐2, early after exposure (Hewett et al., 2000; Iadecola et al., 2001; Pepicelli et al., 2005), suppresses neuronal damage from NMDA but not kainate receptor activation. These data indicate that PG receptor activation is critical to NMDA‐induced neuronal injury. Since PGE2 is the dominant PG secreted under these conditions, the role of specific PG receptors was investigated. Subsequently, others discovered that an antagonist of EP1 (SC51089) administered systemically or intracortically reduces NMDA‐induced neurotoxicity in mice, indicating that EP1 is a downstream neurotoxic effector of NMDA‐induced COX activity (Kawano et al., 2006). These investigators showed in vitro that EP1 enhances neuronal Ca2þ dysregulation induced by NMDA receptor activation through impairment of Naþ–Ca2þ exchange (Kawano et al., 2006). The role of EP2 and EP4 in excitotoxicity appears to be opposite to EP1, namely, that activation of either is neuroprotective. Organotypic cultures from rat hippocampus or spinal cord exposed to excitotoxic challenge show neuroprotection from butaprost, an EP2 agonist (Bilak et al., 2004; Liu et al., 2005). Similarly, in vivo activation of EP4 receptor (which like the EP2 receptor is coupled to Gas) with the agonist ONO‐AE1‐329 is also neuroprotective from glutamate toxicity (Ahmad et al., 2005). Somewhat surprisingly, inhibition of COX‐2 does not abolish the protective effect of EP1 receptor inhibition described earlier (Kawano et al., 2006), as would be predicted if activation of other EP receptors was contributing to the protective effect of EP1 antagonists (Kawano et al., 2006). The relationship between the neuroprotective and neurotoxic effects of the EP receptors following excitotoxic stimulation remains to be clarified.

3.2.2 Aggregated Ab42 Neurotoxicity A major hypothesis concerning AD pathogenesis is that neurodegeneration is mediated, at least in part, by the direct toxic effects of accumulated aggregates of Ab peptides. While precise identification of the neurotoxic species is an area of active investigation, in vitro and in vivo models that produce many forms of aggregated species have been developed and are used widely. For example, aggregated Ab peptides are directly neurotoxic to primary cultures of neurons (just two examples of many are Echeverria et al., 2005; Shie et al., 2005a). One group (Echeverria et al., 2005) reported that pharmacologic activation of EP receptors on wild‐type C57BL/6 mouse primary cerebral cortical neurons using the EP2 agonist butaprost or the EP3/EP4 agonist 1‐hydroxy‐PGE1 suppressed the direct toxicity of low micromolar concentration of aggregated Ab42. Since there are no selective EP2 antagonists, our group (Shie et al., 2005a) used primary neurons derived from EP2
/
mice on the same genetic background or the Balb‐C background and observed no change in the direct neurotoxic effects of aggregated Ab42 in the absence of EP2. It should be kept in mind that Ab peptides are pleiotropic neurotoxins, which not only are directly toxic to neurons but also are indirectly neurotoxic by activating microglia, a process that is strictly dependent on microglial EP2 expression (see later). The relative contribution of direct versus indirect neurotoxicity of Ab peptide in vivo is not yet clear.

3.2.3 Innate Immune Response While the adaptive immune response in the CNS is centrally important in many diseases including ischemic damage, encephalitis, and multiple sclerosis, the innate immune response is now thought to be a major effector in several chronic neurodegenerative diseases such as Alzheimer’s disease (AD), Parkinson’s disease (PD), and amyotrophic lateral sclerosis (ALS), among others. The innate immune response is multifaceted (see earlier), but among its several components cytokine secretion, paracrine oxidative damage to neurons, and phagocytosis of deleterious protein aggregates have been most extensively investigated in brain. As is easy to imagine, some aspects of the innate immune response are damaging to neurons (such as increased free radical stress) whereas others are beneficial (such as clearance of toxic peptide aggregates). While all

E prostanoid receptors in brain physiology and disease

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cells in brain are capable of contributing to the innate immune response, the major contributors are glia and among these microglia are dominant. Cytokine secretion by glia is tightly linked to PG signaling. Two examples from astrocytes are TNF‐a‐ mediated glutamate release and IL‐6 secretion. The former is proposed as a mechanism of astrocyte–neuron synaptic communication, which can be experimentally amplified to neurotoxic levels by the participation of lipopolysaccharide (LPS)‐activated microglia (Bezzi et al., 2001). LPS activation of the microglial innate immune response begins with activation of the co‐receptors CD14 and TLR4, which then initiate a bifurcated signaling cascade with both NFkB‐ and p38‐MAPK‐dependent arms (Imler and Hoffmann, 2001; Akira, 2003). Although discovered for their role in sepsis, these receptors are now known to be part of a larger family of pattern recognition receptors, which respond to both exogenous and endogenous ligands including aggregated Ab peptides and neoantigens expressed by apoptotic cells (Moffatt et al., 1999; Johnson et al., 2003; Fassbender et al., 2004). Thus, like NMDA and excitotoxicity, LPS provides a convenient means to selectively activate a pathologic process via receptors that are similarly activated by other endogenous ligands in disease states. Following intracerebroventricular (ICV) LPS injection, hippocampal CA1 pyramidal neurons undergo a delayed, reversible decrease in dendrite length and spine density without neuron death; spinodendritic degeneration is completely blocked by NSAIDs and in EP2
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mice (Milatovic et al., 2003, 2004). LPS‐ induced inflammatory changes can enhance cognitive decline in aging rodents (Hauss‐Wegrzyniak et al., 2002); reactive oxygen species‐ (ROS‐)induced injury leads to abnormal aging, and impairments in learning‐impaired aged rats are associated with higher levels of oxidative damage than normal aged controls (Nicolle et al., 2001). LPS activation of microglia leads to the elaboration of many cytokines, chemokines, and neuromodulators, a process that can be suppressed in primary cultures of microglia by NSAIDs, suggesting that signaling through some microglial PG receptor is key to this facet of microglial innate immune response (> Table 15‐3). Indeed, the largely responsible receptor appears to be EP2 since its genetic ablation in primary microglia suppresses secretion of over 20 cytokines and chemokines examined (Shie et al., 2005b); the contribution of other EP receptors to this process is not yet known. Thus, EP2 may have opposite effects on neuron survival depending on the pathologic stimulus: in the case of direct damage to neurons from excitotoxicity, EP2 is protective whereas in the case of paracrine damage to neurons from activated microglia, EP2 promotes neurotoxicity. Another aspect of microglial activation that partially overlaps with cytokine and chemokine production is the increased expression and activity of a number of enzymes that generate free radical stress. Microglia themselves are relatively well protected from increased free radical stress through their robust antioxidant defenses (Dringen, 2005). Neurons, in contrast, are relatively vulnerable to free radical stress and, in mixed culture systems as well as in vivo, activation of microglia with LPS leads to paracrine oxidative damage to neurons,which is completely suppressed by NSAIDs (Milatovic et al., 2003; Shie et al., 2005b). Again, the responsible PG receptor appears to be EP2 since its genetic ablation also completely blocks activated microglia‐mediated paracrine damage to wild‐type neurons. Experiments using EP1 antagonists as well as experiments using EP
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mice have failed to show a role for EP1 in activated microglia‐mediated paracrine damage to neurons despite expression of EP1 on microglia (Milatovic et al., 2005; Kawano et al., 2006).

3.2.4 Microglia Phagocytosis of Neurotoxic Peptides As discussed earlier, accumulation of aggregated Ab species is deleterious to neurons through both direct effects and indirect effects that derive from microglial activation. One approach to limit this neurotoxicity is to activate EP4 and possibly EP2 receptors on neurons. Another is to suppress EP2 receptor activity on microglia. A third approach is to enhance clearance of the neurotoxic proteins from brain. Indeed, we have shown that primary cultures of EP2
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microglia on both the Balb‐C and C57BL/6 backgrounds show increased phagocytosis for fluorescently labeled Ab42 (> Table 15‐3). While the primary sequence is the same, higher‐order structure and posttranslational modifications are substantially different between recombinant Ab42 and the Ab species that accumulate in human brain in

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patients with AD. These differences apparently have important consequences. For example, incubation of a slice of human cerebrum from a patient who died with AD (as a source of physiologically aggregated Ab) with wild‐type mouse microglia results in no detectable phagocytosis and clearance of the Ab species; however, opsonization of the Ab species in these experiments leads to extensive clearance of Ab species from human AD tissue (Bard et al., 2000). We have observed that primary mouse microglia from both Balb‐C and C57BL/6 mice that lack EP2 are highly efficient at clearing Ab species from human AD brain slices without prior opsonization (Shie et al., 2005a).

4

Human Brain Diseases and Their Models

4.1 Human Neurologic Diseases Tissue levels of PGs are highly labile due to their rapid metabolism; this confounds interpretation of PG tissue levels in brain, even in rodents where tissue can be immediately flash frozen. For this reason, body fluids are typically used to assess changes in PG production in disease states. Increased PGE2 levels have been demonstrated in cerebrospinal fluid (CSF) from several neurologic diseases (> Table 15‐4).

. Table 15‐4 Increased PGE2 levels in CSF from patients with neurologic disease Disease AD ALS

CJD Ischemic stroke HIV dementia

Increase in CSF PGE2 in neurodegenerative diseases CSF PGE2 levels were highest in very early AD but declined with progressive cognitive impairment fivefold in early AD Sixfold or 2‐ to 10‐fold No increase Sixfold in patients with sporadic or familial forms of CJD as well as variant CJD Twofold during initial 72 h 40% in all HIV‐seropositive patients and positively correlated with degree of cognitive impairment

References Combrinck et al. (2006) Montine et al. (1999) Almer et al. (2002); Ilzecka (2003) Cudkowicz et al. (2006) Minghetti et al. (2000) Minghetti et al. (2002) Aktan et al. (1991) Griffin et al. (1994)

4.2 Disease Models The data summarized in > Table 15‐3 establish that the EP receptors appear to play key roles in several pathophysiologic processes that are proposed to contribute to many neurologic diseases. Furthermore, data from CSF of several degenerative or destructive diseases of brain have shown increased PGE2 in CSF. Combined, these data indicate a potentially critical role for EP receptors in several diseases of the CNS and are the rationale behind several laboratories’ investigations of EP receptors in rodent models of human CNS diseases. However, interpreting the results from complex disease models is often as challenging as studying the actual diseases since multiple pathogenic processes are occurring simultaneously.

4.2.1 Ischemia The pathophysiology of ischemic injury is complex and dependent in part on its severity, the mechanism of ischemia, and the extent of reperfusion. Nevertheless, abundant data support a key role for excitotoxicity and immune‐mediated response to injury in the pathogenesis of brain ischemia. COX‐2 expression is rapidly induced early in neurons in response to massive glutamate release following ischemic injury to brain

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and later in nonneuronal cells, including microglia, astrocytes, and infiltrating mononuclear cells; in the later phases COX‐1 may also contribute significantly (Pepicelli et al., 2005). In rodent models of cerebral ischemia, pharmacologic inhibition of COX‐2 activity and genetic deletion of COX‐2 produce a reduction in infarct size (Nogawa et al., 1997; Nakayama et al., 1998). As mentioned earlier, EP1 receptor null mice or pharmacological inhibition of EP1 signaling rescues brain in a model of transient focal ischemia (Kawano et al., 2006). Given the clear role of excitotoxicity in early damage to neurons in rodent models of ischemic brain damage, the results with EP1 strongly suggest the following sequence: excessive NMDA receptor activation leads to increased COX‐2 activity that leads to increased PGE2 that leads to enhanced activation of neuronal EP1 that is critical to subsequent neuronal injury. Interestingly, reduced infarct size has also been demonstrated in mice lacking EP2 in an in vivo in model of focal cerebral ischemia with or without reperfusion (McCullough et al., 2004; Liu et al., 2005). The similar EP2‐mediated rescue from brain ischemic injury is paradoxical since EP1 and EP2 have opposite effects on excitotoxicity (see earlier). Perhaps this paradox can be explained by the prominent contribution of EP2 to glial pathophysiology, including the innate immune response that is part of ischemic injury to brain.

4.2.2 Alzheimer’s Disease There are several commonly used transgenic mouse models of Familial Alzheimer’s disease (FAD) that derive from overexpression of mutant human amyloid precursor protein (APP) with or without mutant presenilin (PS) 1 proteins. These mice develop age‐dependent amyloid accumulation in concert with significant microglial activation, production of inflammatory cytokines and increased ROS. Treatment with NSAIDs can reduce Ab peptide deposition in vivo in association with reversal of the mild behavioral deficits in these transgenic mice that model the Ab deposition facet of AD (Lim et al., 2000, 2001; Jantzen et al., 2002; Yan et al., 2003). The mechanism by which NSAIDs suppress accumulation of Ab deposits in aged mouse brain is likely attributed to their inhibition of COX isozymes. An alternative hypothesis advanced by recent in vitro studies asserts that high concentrations of some but not all NSAIDs can alter g‐secretase activity and reduce the ratio of Ab 42 to 38 in cultured cells (Weggen et al., 2001, 2003a, b); however, pharmacologically relevant doses of NSAIDs in vivo challenge this proposal (Eriksen et al., 2003; Lanz et al., 2005). In aged APPSwe‐PS1DE9 mice, deletion of the EP2 receptor is also associated with lower levels of total Ab40 and Ab42 peptides and fewer accumulated amyloid deposits (Liang et al., 2005). The deletion of the EP2 receptor in these transgenic mice also results in marked decreases in lipid peroxidation, with significant reductions in neuronal lipid peroxidation (Liang et al., 2005) that parallel what is seen in the more direct LPS model described earlier (Montine et al., 2002; Milatovic et al., 2004). One interpretation of these data is that the neuronal oxidative damage observed in this model is secondary to Ab activation of microglial CD14‐dependent signaling (Fassbender et al., 2004). Emerging data suggest a reinforcing cycle between oxidative damage and formation of neurotoxic Ab aggregates (Woltjer et al., 2007), raising the question of which comes first in vivo. Data from these same mouse models suggests that oxidative damage may precede and possibly trigger Ab deposition (Pratico et al., 2001). Another interpretation is that accumulation of Ab deposits are decreased in the brains of these mice that lack EP2 because of enhanced microglial phagocytosis of Ab species. Whether lack of EP2 leads to reduced Ab deposition because of decreased microglial CD14‐ mediated paracrine oxidative damage to neurons or because of increased phagocytosis is not clear. Regardless, these data underscore a highly desirable dual phenotype from ablation of microglial EP2 signaling in these transgenic mouse models of FAD.

4.2.3 Parkinson’s Disease PD is another age‐related neurodegenerative disease that prominently involves degeneration of the dopaminergic neurons of the substantia nigra pars compacta. As in AD, a major role for neuroinflammation is now proposed in PD as evidenced by significant microglial activation with complement activation in

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autopsy specimens (reviewed in McGeer and McGeer, 2004). Significantly, COX‐2 and downstream PG signaling appear to play a role in disease progression in rodent models of PD employing the selective dopaminergic neurotoxin 1‐methyl‐4‐phenyl‐1,2,3,6‐tetrahydropyridine (MPTP). In these models, inhibition of COX‐2, either using genetic COX‐2 knockout strategies or COX‐2 inhibitors, partially protects against dopaminergic neurodegeneration and development of motor symptoms (Feng et al., 2002; Teismann et al., 2003). In strict analogy to our previous experiments with AD brain, we have recently shown that ablation of EP2 also significantly enhanced microglia‐mediated ex vivo clearance of a‐synuclein aggregates from the mesocortex of Lewy body disease patients while significantly attenuating neurotoxicity and the extent of a‐synuclein aggregation in mice intoxicated with MPTP (Jin et al., 2007).

4.2.4 Amyotrophic Lateral Sclerosis ALS is characterized by the progressive loss of motor neurons in ventral spinal cord and motor cortex. Pathologic findings in ALS suggest an inflammatory component to this disease involving activated microglia, astrocytes (McGeer and McGeer, 2002), and increased COX‐2 expression (Yasojima et al., 2001; Maihofner et al., 2003; Yiangou et al., 2006). Indeed, some have shown neuroprotection in an organotypic model of ALS by suppressing COX‐2 activity (Drachman and Rothstein, 2000). An extensively characterized familial form of ALS is caused by mutations in the copper/zinc superoxide dismutase (SOD1) gene (Gurney et al., 1994; Wong et al., 1995; Bruijn et al., 1998). Transgenic mice that express mutant forms of SOD1 recapitulate the human disease and develop paralysis associated with loss of motor neurons and premature death. Levels of COX‐2 mRNA and PGE2 are increased in mutant SOD mice (Almer et al., 2001; Klivenyi et al., 2004) and administration of COX‐2 inhibitors beginning before onset of hindlimb paralysis reduces levels of CSF PGE2 and extends survival (Drachman et al., 2002; Pompl et al., 2003; Klivenyi et al., 2004). These observations have suggested that COX‐2, via its downstream PG products, can promote motor neuron injury in this model. Investigations of the contribution of specific EP receptors in this model are underway.

5

Conclusion

The dichotomous neuroprotective and neurotoxic role of prostaglandin receptor activation in human neurological disease is only partially resolved. Studies to elucidate the balance between neuronal cell death and survival and its relationship to coordinated and differential EP receptor activation are ongoing and have the potential not only to enlighten scientists and clinicians to neurodegenerative disease pathophysiology, but to guide us to develop efficacious therapies that selectively inhibit neurotoxicity while potentiating neuroprotection mediated by neuroinflammatory pathways.

References Ahmad AS, Ahmad M, de Brum‐Fernandes AJ, Dore S. 2005. Prostaglandin EP4 receptor agonist protects against acute neurotoxicity. Brain Res 1066: 71-77. Akaneya Y, Tsumoto T. 2006. Bidirectional trafficking of prostaglandin E2 receptors involved in long‐term potentiation in visual cortex. J Neurosci 26: 10209-10221. Akaogi J, Yamada H, Kuroda Y, Nacionales DC, Reeves WH, et al. 2004. Prostaglandin E2 receptors EP2 and EP4 are up‐regulated in peritoneal macrophages and joints of pristane‐treated mice and

modulate TNF‐alpha and IL‐6 production. J Leukoc Biol 76: 227-236. Akira S. 2003. Toll‐like receptor signaling. J Biol Chem 278: 38105-38108. Aktan S, Aykut C, Ercan S. 1991. Leukotriene C4 and prostaglandin E2 activities in the serum and cerebrospinal fluid during acute cerebral ischemia. Prostaglandins Leukot Essent Fatty Acids 43: 247-249. Almer G, Guegan C, Teismann P, Naini A, Rosoklija G, et al. 2001. Increased expression of the pro‐inflammatory

E prostanoid receptors in brain physiology and disease enzyme cyclooxygenase‐2 in amyotrophic lateral sclerosis. Ann Neurol 49: 176-185. Almer G, Teismann P, Stevic Z, Halaschek‐Wiener J, Deecke L, et al. 2002. Increased levels of the pro‐inflammatory prostaglandin PGE2 in CSF from ALS patients. Neurology 58: 1277-1279. Aronoff DM, Canetti C, Peters‐Golden M. 2004. Prostaglandin E2 inhibits alveolar macrophage phagocytosis through an E‐prostanoid 2 receptor‐mediated increase in intracellular cyclic AMP. J Immunol 173: 559-565. Baratelli FE, Heuze‐Vourc’h N, Krysan K, Dohadwala M, Riedl K, et al. 2004. Prostaglandin E2‐dependent enhancement of tissue inhibitors of metalloproteinases‐1 production limits dendritic cell migration through extracellular matrix. J Immunol 173: 5458-5466. Bard F, Cannon C, Barbour R, Burke RL, Games D, et al. 2000. Peripherally administered antibodies against amyloid beta‐ peptide enter the central nervous system and reduce pathology in a mouse model of Alzheimer disease. Nat Med 6: 916-919. Batshake B, Nilsson C, Sundelin J. 1995. Molecular characterization of the mouse prostanoid EP1 receptor gene. Eur J Biochem 231: 809-814. Bezzi P, Carmignoto G, Pasti L, Vesce S, Rossi D, et al. 1998. Prostaglandins stimulate calcium dependent glutamate release in astrocytes. Nature 391: 281-285. Bezzi P, Domercq M, Brambilla L, Galli R, Schols D, et al. 2001. CXCR4‐activated astrocyte glutamate release via TNFalpha: Amplification by microglia triggers neurotoxicity. Nat Neurosci 4: 702-710. Bhattacharya M, Peri K, Almazan G, Ribeiro‐da‐Silva A, Shichi H, et al. 1998. Nuclear localization of prostaglandin E2 receptors. Proc Natl Acad Sci USA 95: 15792-15797. Bilak M, Wu L, Wang Q, Haughey N, Conant K, et al. 2004. PGE2 receptors rescue motor neurons in a model of amyotrophic lateral sclerosis. Ann Neurol 56: 240-248. Bjornson AB, Knippenberg RW, Bjornson HS. 1988. Nonsteroidal anti‐inflammatory drugs correct the bactericidal defect of polymorphonuclear leukocytes in a guinea pig model of thermal injury. J Infect Dis 157: 959-967. Borda ES, Tenenbaum A, Sales ME, Rumi L, Sterin‐Borda L. 1998. Role of arachidonic acid metabolites in the action of a beta adrenergic agonist on human monocyte phagocytosis. Prostaglandins Leukot Essent Fatty Acids 58: 85-90. Boutaud O, Ou JJ, Chaurand P, Caprioli RM, Montine TJ, et al. 2002. Prostaglandin H2 (PGH2) accelerates formation of amyloid beta1‐42 oligomers. J Neurochem 82: 1003-1006. Breyer MD, Breyer RM. 2001. G protein‐coupled prostanoid receptors and the kidney. Annu Rev Physiol 63: 579-605. Breyer RM, Bagdassarian CK, Myers SA, Breyer MD. 2001. Prostanoid receptors: Subtypes and signaling. Ann Rev Pharmacol Toxicol 41: 661-690.

15

Bruijn LI, Houseweart MK, Kato S, Anderson KL, Anderson SD, et al. 1998. Aggregation and motor neuron toxicity of an ALS‐linked SOD1 mutant independent from wild‐type SOD1. Science 281: 1851-1854. Caggiano AO, Kraig RP. 1999. Prostaglandin E receptor subtypes in cultured rat microglia and their role in reducing lipopolysaccharide‐induced interleukin‐1beta production. J Neurochem 72: 565-575. Campbell WB, Halushka PV. 2001. Lipid‐derived autacoids. Goodman & Gilman’s the Pharmacological Basis of Therapeutics, 9th Edition. Hardman JG, Limbird LE, editors. New York: McGraw‐Hill; pp. 601-616. Candelario‐Jalil E, Slawik H, Ridelis I, Waschbisch A, Akundi RS, et al. 2005. Regional distribution of the prostaglandin E2 receptor EP1 in the rat brain: Accumulation in Purkinje cells of the cerebellum. J Mol Neurosci 27: 303-310. Canning BJ, Hmieleski RR, Spannhake EW, Jakab GJ. 1991. Ozone reduces murine alveolar and peritoneal macrophage phagocytosis: The role of prostanoids. Am J Physiol 261: L277-L282. Choi JS, Kim HY, Chun MH, Chung JW, Lee MY. 2006. Expression of prostaglandin E(2) receptor subtypes, EP2 and EP4, in the rat hippocampus after cerebral ischemia and ischemic tolerance. Cell Tissue Res 324: 203-211. Combrinck M, Williams J, De Berardinis MA, Warden D, Puopolo M, et al. 2006. Levels of CSF prostaglandin E2, cognitive decline, and survival in Alzheimer’s disease. J Neurol Neurosurg Psychiatry 77: 85-88. Cudkowicz ME, Shefner JM, Schoenfeld DA, Zhang H, Andreasson KI, et al. 2006. Trial of celecoxib in amyotrophic lateral sclerosis. Ann Neurol 60: 22-31. Davis RJ, Murdoch CE, Ali M, Purbrick S, Ravid R, et al. 2004. EP4 prostanoid receptor‐mediated vasodilatation of human middle cerebral arteries. Br J Pharmacol 141: 580-585. Difranco E, Subbanagounder G, Kim S, Murthi K, Taneda S, et al. 1994. Formation and stability of pyrrole adducts in the reaction of levuglandin E2 with proteins. Chem Res Toxicol 8: 61-67. Drachman DB, Rothstein JD. 2000. Inhibition of cyclooxygenase‐2 protects motor neurons in an organotypic model of amyotrophic lateral sclerosis. Ann Neurol 48: 792-795. Drachman DB, Frank K, Dykes‐Hoberg M, Teismann P, Almer G, et al. 2002. Cyclooxygenase 2 inhibition protects motor neurons and prolongs survival in a transgenic mouse model of ALS. Ann Neurol 52: 771-778. Dringen R. 2005. Oxidative and antioxidative potential of brain microglial cells. Antioxid Redox Signal 7: 1223-1233. Echeverria V, Clerman A, Dore S. 2005. Stimulation of PGE receptors EP2 and EP4 protects cultured neurons against oxidative stress and cell death following beta‐amyloid exposure. Eur J Neurosci 22: 2199-2206.

397

398

15

E prostanoid receptors in brain physiology and disease

Ek M, Arias C, Sawchenko P, Ericsson‐Dahlstrand A. 2000. Distribution of the EP3 prostaglandin E2 receptor subtype in the rat brain: Relationship to sites of interleukin‐1‐ induced cellular responsiveness. J Comp Neurol 428: 5-20. Engblom D, Ek M, Ericsson‐Dahlstrand A, Blomqvist A. 2004. EP3 and EP4 receptor mRNA expression in peptidergic cell groups of the rat parabrachial nucleus. Neuroscience 126: 989-999. Eriksen JL, Sagi SA, Smith TE, Weggen S, Das P, et al. 2003. NSAIDs and enantiomers of flurbiprofen target gamma‐ secretase and lower Abeta 42 in vivo. J Clin Invest 112: 440-449. Fassbender K, Walter S, Kuhl S, Landmann R, Ishii K, et al. 2004. The LPS receptor (CD14) links innate immunity with Alzheimer’s disease. FASEB J 18: 203-205. Fedyk ER, Phipps RP. 1996. Prostaglandin E2 receptors of the EP2 and EP4 subtypes regulate activation and differentiation of mouse B lymphocytes to IgE‐secreting cells. Proc Natl Acad Sci USA 93: 10978-10983. Feng ZH, Wang TG, Li DD, Fung P, Wilson BC, et al. 2002. Cyclooxygenase‐2‐deficient mice are resistant to 1‐methyl‐ 4‐phenyl1, 2,3,6‐tetrahydropyridine‐induced damage of dopaminergic neurons in the substantia nigra. Neurosci Lett 329: 354-358. Fennekohl A, Sugimoto Y, Segi E, Maruyama T, Ichikawa A, et al. 2002. Contribution of the two Gs‐coupled PGE2‐ receptors EP2‐receptor and EP4‐receptor to the inhibition by PGE2 of the LPS‐induced TNFalpha‐formation in Kupffer cells from EP2‐or EP4‐receptor‐deficient mice. Pivotal role for the EP4‐receptor in wild type Kupffer cells. J Hepatol 36: 328-334. Fiebich BL, Schleicher S, Spleiss O, Czygan M, Hull M. 2001. Mechanisms of prostaglandin E2‐induced interleukin‐6 release in astrocytes: Possible involvement of EP4‐like receptors, p38 mitogen‐activated protein kinase and protein kinase C. J Neurochem 79: 950-958. Gardiner PJ. 1986. Characterization of prostanoid relaxant/ inhibitory receptors (psi) using a highly selective agonist, TR4979. Br J Pharmacol 87: 45-56. Gilmour MI, Park P, Doerfler D, Selgrade MK. 1993. Factors that influence the suppression of pulmonary antibacterial defenses in mice exposed to ozone. Exp Lung Res 19: 299-314. Griffin DE, Wesselingh SL, McArthur JC. 1994. Elevated central nervous system prostaglandins in human immunodeficiency virus‐associated dementia. Ann Neurol 35: 592-597. Gurer US, Palanduz A, Gurbuz B, Yildirmak Y, Cevikbas A, et al. 2002. Effect of antipyretics on polymorphonuclear leukocyte functions in children. Int Immunopharmacol 2: 1599-1602. Gurney ME, Pu H, Chiu AY, Dal Canto MC, Polchow CY, et al. 1994. Motor neuron degeneration in mice that express a

human Cu,Zn superoxide dismutase mutation. Science 264: 1772-1775. Harizi H, Grosset C, Gualde N. 2003. Prostaglandin E2 modulates dendritic cell function via EP2 and EP4 receptor subtypes. J Leukoc Biol 73: 756-763. Hata AN, Breyer RM. 2004. Pharmacology and signaling of prostaglandin receptors: Multiple roles in inflammation and immune modulation. Pharmacol Ther 103: 147-166. Hauss‐Wegrzyniak B, Lynch MA, Vraniak PD, Wenk GL. 2002. Chronic brain inflammation results in cell loss in the entorhinal cortex and impaired LTP in perforant path‐granule cell synapses. Exp Neurol 176: 336-341. Hewett SJ, Uliasz TF, Vidwans AS, Hewett JA. 2000. Cyclooxygenase‐2 contributes to N‐methyl‐D‐aspartate‐ mediated neuronal cell death in primary cortical cell culture. J Pharmacol Exp Ther 293: 417-425. Hirai H, Tanaka K, Yoshie O, Ogawa K, Kenmotsu K, et al. 2001. Prostaglandin D2 selectively induces chemotaxis in T helper type 2 cells, eosinophils, and basophils via seven‐ transmembrane receptor CRTH2. J Exp Med 193: 255-261. Hubbard NE, Lee S, Lim D, Erickson KL. 2001. Differential mRNA expression of prostaglandin receptor subtypes in macrophage activation. Prostaglandins Leukot Essent Fatty Acids 65: 287-294. Hutchison DL, Myers RL. 1987. Prostaglandin‐mediated suppression of macrophage phagocytosis of Listeria monocytogenes. Cell Immunol 110: 68-76. Iadecola C, Niwa K, Nogawa S, Zhao X, Nagayama M, et al. 2001. Reduced susceptibility to ischemic brain injury and N‐methyl‐D‐aspartate‐mediated neurotoxicity in cyclooxygenase‐2‐deficient mice. Proc Natl Acad Sci USA 98: 1294-1299. Ilzecka J. 2003. Prostaglandin E2 is increased in amyotrophic lateral sclerosis patients. Acta Neurol Scand 108: 125-129. Imler JL, Hoffmann JA. 2001. Toll receptors in innate immunity. Trends Cell Biol 11: 304-311. Jantzen PT, Connor KE, DiCarlo G, Wenk G, Wallace J, et al. 2002. Microglial activation and b‐amyloid deposit reduction caused by a nitric oxide‐releasing nonsteroidal anti‐ inflammatory drug in amyloid precursor protein plus presenilin‐1 transgenic mice. J Neurosci 22: 2246-2254. Jin J, Shie FS, Liu J, Wang Y, Davis J, et al. 2007. Prostaglandin E2 receptor subtype 2 (EP2) regulates microglial activation and associated neurotoxicity induced by aggregated alpha‐ synuclein. J Neuroinflammation 4: 2. Johnson GB, Brunn GJ, Platt JL. 2003. Activation of mammalian toll‐like receptors by endogenous agonists. Crit Rev Immunol 23: 15-44. Kabashima K, Murata T, Tanaka H, Matsuoka T, Sakata D, et al. 2003. Thromboxane A2 modulates interaction of dendritic cells and T cells and regulates acquired immunity. Nat Immunol 4: 694-701.

E prostanoid receptors in brain physiology and disease Kaufmann WE, Andreasson KI, Isakson PC, Worley PF. 1997. Cyclooxygenases and the central nervous system. Prostaglandins 54: 601-624. Kawano T, Anrather J, Zhou P, Park L, Wang G, et al. 2006. Prostaglandin E2 EP1 receptors: Downstream effectors of COX‐2 neurotoxicity. Nat Med 12: 225-229. Kennedy CR, Zhang Y, Brandon S, Guan Y, Coffee K, et al. 1999. Salt‐sensitive hypertension and reduced fertility in mice lacking the prostaglandin EP2 receptor. Nat Med 5: 217-220. Kitanaka J, Hashimoto H, Gotoh M, Kondo K, Sakata K, et al. 1996. Expression pattern of messenger RNA’s for prostanoid receptors in glial cell cultures. Brain Res 707: 282-287. Klivenyi P, Kiaei M, Gardian G, Calingasan NY, Beal MF. 2004. Additive neuroprotective effects of creatine and cyclooxygenase 2 inhibitors in a transgenic mouse model of amyotrophic lateral sclerosis. J Neurochem 88: 576-582. Kudo I, Murakami M. 2002. Phospholipase A2 enzymes. Prostaglandins Other Lipid Mediat 68‐69: 3-58. Lanz TA, Fici GJ, Merchant KM. 2005. Lack of specific amyloid‐{beta}(1–42) suppression by nonsteroidal anti‐ inflammatory drugs in young, plaque‐free Tg2576 mice and in guinea pig neuronal cultures. J Pharmacol Exp Ther 312: 399-406. Largo R, Diez‐Ortego I, Sanchez‐Pernaute O, Lopez‐ Armada MJ, Alvarez‐Soria MA, et al. 2004. EP2/EP4 signalling inhibits monocyte chemoattractant protein‐1 production induced by interleukin 1beta in synovial fibroblasts. Ann Rheum Dis 63: 1197-1204. Liang X, Wang Q, Hand T, Wu L, Breyer RM, et al. 2005. Deletion of the prostaglandin E2 EP2 receptor reduces oxidative damage and amyloid burden in a model of Alzheimer’s disease. J Neurosci 25: 10180-10187. Lim GP, Yang F, Chu T, Chen P, Beech W, et al. 2000. Ibuprofen suppresses plaque pathology and inflammation in a mouse model for Alzheimer’s disease. J Neurosci 20: 5709-5714. Lim GP, Yang F, Chu T, Gahtan E, Ubeda O, et al. 2001. Ibuprofen effects on Alzheimer pathology and open field activity in APPsw transgenic mice. Neurobiol Aging 22: 983-991. Liu D, Wu L, Breyer R, Mattson MP, Andreasson K. 2005. Neuroprotection by the PGE2 EP2 receptor in permanent focal cerebral ischemia. Ann Neurol 57: 758-761. Maihofner C, Probst‐Cousin S, Bergmann M, Neuhuber W, Neundorfer B, et al. 2003. Expression and localization of cyclooxygenase‐1 and ‐2 in human sporadic amyotrophic lateral sclerosis. Eur J Neurosci 18: 1527-1534. McCullough L, Wu L, Haughey N, Liang X, Hand T, et al. 2004. Neuroprotective function of the PGE2 EP2 receptor in cerebral ischemia. J Neurosci 24: 257-268.

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McGeer PL, McGeer EG. 2002. Inflammatory processes in amyotrophic lateral sclerosis. Muscle Nerve 26: 459-470. McGeer PL, McGeer EG. 2004. Inflammation and neurodegeneration in Parkinson’s disease. Parkinsonism Relat Disord 10 Suppl 1: S3-S7. Milatovic D, Zaja‐Milatovic S, Montine KS, Horner PJ, Montine TJ. 2003. Pharmacologic suppression of neuronal oxidative damage and dendritic degeneration following direct activation of glial innate immunity in mouse cerebrum. J Neurochem 87: 1518-1526. Milatovic D, Zaja‐Milatovic S, Montine KS, Shie FS, Montine TJ. 2004. Neuronal oxidative damage and dendritic degeneration following activation of CD14‐dependent innate immune response in vivo. J Neuroinflammation 1: 20. Milatovic D, Zaja‐Milatovic S, Montine KS, Nivison M, Montine TJ. 2005. CD14‐dependent innate immunity‐ mediated neuronal damage in vivo is suppressed by NSAIDS and ablation of a prostaglandin E2 receptor, EP2. Curr Med Chem 5: 151-156. Minami T, Nakano H, Kobayashi T, Sugimoto Y, Ushikubi F, et al. 2001. Characterization of EP receptor subtypes responsible for prostaglandin E2‐induced pain responses by use of EP1 and EP3 receptor knockout mice. Br J Pharmacol 133: 438-444. Minghetti L, Nicolini A, Polazzi E, Creminon C, Maclouf J, et al. 1997. Prostaglandin E2 downregulates inducible nitric oxide synthase expression in microglia by increasing cAMP levels. Adv Exp Med Biol 433: 181-184. Minghetti L, Greco A, Cardone F, Puopolo M, Ladogana A, et al. 2000. Increased brain synthesis of prostaglandin E2 and F2‐isoprostane in human and experimental transmissible spongiform encephalopathies. J Neuropathol Exp Neurol 59: 866-871. Minghetti L, Cardone F, Greco A, Puopolo M, Levi G, et al. 2002. Increased CSF levels of prostaglandin E(2) in variant Creutzfeldt‐Jakob disease. Neurology 58: 127-129. Moffatt OD, Devitt A, Bell ED, Simmons DL, Gregory CD. 1999. Macrophage recognition of ICAM‐3 on apoptotic leukocytes. J Immunol 162: 6800-6810. Montine TJ, Sidell KR, Crews BC, Markesbery WR, Marnett LJ, et al. 1999. Elevated CSF prostaglandin E2 levels in patients with probable AD. Neurology 53: 1495-1498. Montine TJ, Milatovic D, Gupta RC, Valyi‐Nagy T, Morrow JD, et al. 2002. Neuronal oxidative damage from activated innate immunity is EP2 receptor‐dependent. J Neurochem 83: 463-470. Nakamura K, Kaneko T, Yamashita Y, Hasegawa H, Katoh H, et al. 2000. Immunohistochemical localization of prostaglandin EP3 receptor in the rat central nervous system. J Comp Neurol 421: 543-569. Nakamura K, Li YQ, Kaneko T, Katoh H, Negishi M. 2001. Prostaglandin EP3 receptor protein in serotonin and

399

400

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catecholamine cell groups: A double immunofluorescence study in the rat brain. Neuroscience 103: 763-775. Nakayama M, Uchimura K, Zhu RL, Nagayama T, Rose ME, et al. 1998. Cyclooxygenase‐2 inhibition prevents delayed death of CA1 hippocampal neurons following global ischemia. Proc Natl Acad Sci USA 95: 10954-10959. Nakayama Y, Omote K, Namiki A. 2002. Role of prostaglandin receptor EP1 in the spinal dorsal horn in carrageenan‐ induced inflammatory pain. Anesthesiology 97: 1254-1262. Nataraj C, Thomas DW, Tilley SL, Nguyen MT, Mannon R, et al. 2001. Receptors for prostaglandin E(2) that regulate cellular immune responses in the mouse. J Clin Invest 108: 1229-1235. Negishi M, Sugimoto Y, Irie A, Narumiya S, Ichikawa A. 1993. Two isoforms of prostaglandin E receptor EP3 subtype. Different COOH‐terminal domains determine sensitivity to agonist‐induced desensitization. J Biol Chem 268: 9517-9521. Nguyen M, Solle M, Audoly LP, Tilley SL, Stock JL, et al. 2002. Receptors and signaling mechanisms required for prostaglandin E2‐mediated regulation of mast cell degranulation and IL‐6 production. J Immunol 169: 4586-4593. Nicolle MM, Gonzalez J, Sugaya K, Baskerville KA, Bryan D, et al. 2001. Signatures of hippocampal oxidative stress in aged spatial learning‐impaired rodents. Neuroscience 107: 415-431. Nogawa S, Zhang F, Ross ME, Iadecola C. 1997. Cyclo‐ oxygenase‐2 gene expression in neurons contributes to ischemic brain damage. J Neurosci 17: 2746-2755. Noguchi K, Iwasaki K, Shitashige M, Umeda M, Izumi Y, et al. 2001. Downregulation of lipopolysaccharide‐induced intercellular adhesion molecule‐1 expression via EP2/EP4 receptors by prostaglandin E2 in human fibroblasts. Inflammation 25: 75-81. Oka T, Oka K, Schammell TE, Lee C, Kelly JF, et al. 2000. Relationship of EP1‐4 prostaglandin receptors with rat hypothalamic cell groups involved in lipopolysaccharide fever responses. J Comp Neurol 428: 20-32. Pepicelli O, Fedele E, Berardi M, Raiteri M, Levi G, et al. 2005. Cyclo‐oxygenase‐1 and ‐2 differently contribute to prostaglandin E2 synthesis and lipid peroxidation after in vivo activation of N‐methyl‐D‐aspartate receptors in rat hippocampus. J Neurochem 93: 1561-1567. Planells‐Cases R, Lerma J, Ferrer‐Montiel A. 2006. Pharmacological intervention at ionotropic glutamate receptor complexes. Curr Pharm Des 12: 3583-3596. Pompl PN, Ho L, Bianchi M, McManus T, Qin W, et al. 2003. A therapeutic role for cyclooxygenase‐2 inhibitors in a transgenic mouse model of amyotrophic lateral sclerosis. FASEB J 17: 725-727. Pratico D, Uryu K, Leight S, Trojanowski JQ, Lee VM. 2001. Increased lipid peroxidation precedes amyloid plaque

formation in an animal model of Alzheimer amyloidosis. J Neurosci 21: 4183-4187. Shie FS, Breyer RM, Montine TJ. 2005a. Microglia lacking E prostanoid receptor subtype 2 have enhanced A{beta} phagocytosis yet lack A{beta}‐activated neurotoxicity. Am J Pathol 166: 1163-1172. Shie FS, Montine KS, Breyer RM, Montine TJ. 2005b. Microglial EP2 is critical to neurotoxicity from activated cerebral innate immunity. Glia 52: 70-77. Siedlik PH, Marnett LJ. 1984. Oxidizing radical generation by prostaglandin H synthase. Meth Enzymol 105: 412-416. Slawik H, Volk B, Fiebich B, Hull M. 2004. Microglial expression of prostaglandin EP3 receptor in excitotoxic lesions in the rat striatum. Neurochem Int 45: 653-660. Stock JL, Shinjo K, Burkhardt J, Roach M, Taniguchi K, et al. 2001. The prostaglandin E2 EP1 receptor mediates pain perception and regulates blood pressure. J Clin Invest 107: 325-331. Sugimoto Y, Shigemoto R, Namba T, Negishi M, Mizuno N, et al. 1994. Distribution of the messenger RNA for the prostaglandin E receptor subtype EP3 in the mouse nervous system. Neuroscience 62: 919-928. Teismann P, Tieu K, Choi DK, Wu DC, Naini A, et al. 2003. Cyclooxygenase‐2 is instrumental in Parkinson’s disease neurodegeneration. Proc Natl Acad Sci USA 100: 5473-5478. Treffkorn L, Scheibe R, Maruyama T, Dieter P. 2004. PGE2 exerts its effect on the LPS‐induced release of TNF‐alpha, ET‐1, IL‐1alpha, IL‐6 and IL‐10 via the EP2 and EP4 receptor in rat liver macrophages. Prostaglandins Other Lipid Mediat 74: 113-123. Vassiliou E, Jing H, Ganea D. 2003. Prostaglandin E2 inhibits TNF production in murine bone marrow‐derived dendritic cells. Cell Immunol 223: 120-132. Walch L, de Montpreville V, Brink C, Norel X. 2001. Prostanoid EP(1)‐ and TP‐receptors involved in the contraction of human pulmonary veins. Br J Pharmacol 134: 1671-1678. Warner TD, Giuliano F, Vojnovic I, Bukasa A, Mitchell JA, et al. 1999. Nonsteroid drug selectivities for cyclo‐ oxygenase‐1 rather than cyclo‐oxygenase‐2 are associated with human gastrointestinal toxicity: A full in vitro analysis. Proc Natl Acad Sci USA 96: 7563-7568. Waschbisch A, Fiebich BL, Akundi RS, Schmitz ML, Hoozemans JJ, et al. 2006. Interleukin‐1 beta‐induced expression of the prostaglandin E‐receptor subtype EP3 in U373 astrocytoma cells depends on protein kinase C and nuclear factor‐kappa B. J Neurochem 96: 680-693. Weggen S, Eriksen JL, Das P, Sagi SA, Wang R, et al. 2001. A subset of NSAIDs lower amyloidogenic Abeta42 independently of cyclooxygenase activity. Nature 414: 212-216. Weggen S, Eriksen JL, Sagi SA, Pietrzik CU, Golde TE, et al. 2003a. Abeta42‐lowering nonsteroidal anti‐inflammatory

E prostanoid receptors in brain physiology and disease drugs preserve intramembrane cleavage of the amyloid precursor protein (APP) and ErbB‐4 receptor and signaling through the APP intracellular domain. J Biol Chem 278: 30748-30754. Weggen S, Eriksen JL, Sagi SA, Pietrzik CU, Ozols V, et al. 2003b. Evidence that nonsteroidal anti‐inflammatory drugs decrease amyloid beta 42 production by direct modulation of gamma‐secretase activity. J Biol Chem 278: 31831-31837. Woltjer RL, McMahan W, Milatovic D, Kjerulf JD, Shie FS, et al. 2007. Effects of chemical chaperones on oxidative stress and detergent‐insoluble species formation following conditional expression of amyloid precursor protein carboxy‐terminal fragment. Neurobiol Dis 25: 427-437. Wong PC, Pardo CA, Borchelt DR, Lee MK, Copeland NG, et al. 1995. An adverse property of a familial ALS‐linked SOD1 mutation causes motor neuron disease characterized by vacuolar degeneration of mitochondria. Neuron 14: 1105-1116. Woo SK, Roszkowski P, Waterbury LD, Garay GL. 1986. Gastric mucosal binding studies with enprostil: A potent anti‐ulcer prostaglandin. Prostaglandins 32: 243-257. Yamagata K, Andreasson KI, Kaufmann WE, Barnes CA, Worley PF. 1993. Expression of a mitogen‐inducible

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cyclooxygenase in brain neurons: Regulation by synaptic activity and glucocorticoids. Neuron 11: 371-386. Yan Q, Zhang J, Liu H, Babu‐Khan S, Vassar R, et al. 2003. Anti‐inflammatory drug therapy alters beta‐amyloid processing and deposition in an animal model of Alzheimer’s disease. J Neurosci 23: 7504-7509. Yang L, Yamagata N, Yadav R, Brandon S, Courtney RL, et al. 2003. Cancer‐associated immunodeficiency and dendritic cell abnormalities mediated by the prostaglandin EP2 receptor. J Clin Invest 111: 727-735. Yasojima K, Tourtellotte WW, McGeer EG, McGeer PL. 2001. Marked increase in cyclooxygenase‐2 in ALS spinal cord: Implications for therapy. Neurology 57: 952-956. Yiangou Y, Facer P, Durrenberger P, Chessell IP, Naylor A, et al. 2006. COX‐2, CB2 and P2X7‐immunoreactivities are increased in activated microglial cells/macrophages of multiple sclerosis and amyotrophic lateral sclerosis spinal cord. BMC Neurol 6: 12. Zhang J, Rivest S. 1999. Distribution, regulation and colocalization of the genes encoding the EP2‐ and EP4‐PGE2 receptors in the rat brain and neuronal responses to systemic inflammation. Eur J Neurosci 11: 2651-2668.

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Nitric Oxide and other Diffusible Messengers

J. P. Kiss

1 1.1 1.1.1 1.1.2 1.1.3 1.1.4 1.1.5 1.2 1.3 1.3.1 1.3.2 1.3.3 1.4 1.5

Nitric Oxide . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 404 Synthesis of NO, Properties of NOS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 404 Synthesis of NO . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 404 Regulation of NOS Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 406 Subcellular Localization of NOS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 406 Expression of NOS Splice Variants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 407 NOS Inhibitors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 407 Effector Mechanisms of NO . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 407 NO and nNOS in the Central Nervous System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 408 General Features . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 408 NO and LTP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 408 NO as a Nonsynaptic Link between Glutamatergic and Monoaminergic Neurons . . . . . . . . . . . . 409 NO and Neurotoxicity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 410 NO in the Peripheral Nervous System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 411

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Carbon Monoxide . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 412

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Hydrogen Peroxide . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 412

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2008 Springer ScienceþBusiness Media, LLC.

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Abstract: The era of diffusible messengers has begun in the late 1980s when a revolutionary discovery (Palmer et al., 1987) clarified that the previously described, chemically unknown compound, endothelium derived relaxing factor (EDRF) is identical with nitric oxide (NO). The scientific community was astonished by the fact that such a simple gaseous molecule as NO can play a very important physiological role in the regulation of the resistance of blood vessels. The research of nitrergic systems and processes has started with huge enthusiasm and the number of NO‐related research papers increased exponentially. NO has been elected to the ‘‘Molecule of the Year’’ by the prestigious journal Science in 1992, and the Nobel Prize in Medicine was given to Furchgott, Ignarro and Murad ‘‘for their discoveries concerning NO as a signalling molecule in the cardiovascular system’’ in 1998, only 11 years after the original finding. The results of the last two decades revealed that NO has very important functions not only in the cardiovascular but also in the immune and nervous system. This chapter will focus primarily on the role of NO in the central and peripheral nervous system, but occasionally other areas will also be covered. In addition, other recently recognized diffusible compounds with possible neuromodulator function will also be briefly discussed. List of Abbreviations: CAPON, carboxy‐terminal PDZ ligand of nNOS; CNS, Central nervous system; EDRF, endothelium derived relaxing factor; L‐NAME, N(omega)‐nitro‐L‐arginine methyl ester; LTP, long‐term potentiation; NANC, non‐adrenerg non‐cholinerg; NO, nitric oxide; NOS, nitric oxide synthase; PARS, polyADP‐ribose synthase; PIN, protein inhibitor of nNOS; RNS, reactive nitrogen species; ROS, reactive oxygen species; sGC, soluble guanylyl cyclase; SOD, superoxide dismutase; THB, tetrahydrobiopterin

1

Nitric Oxide

Nitric oxide (NO) is a simple gas molecule with an unpaired electron that makes this compound a very reactive free radical. Because of its physicochemical properties NO is a highly diffusible gas that crosses the biological membranes without any difficulty. Although its half‐life is only a few seconds even during this short period it can diffuse away a few hundred mm from the site of production.

1.1 Synthesis of NO, Properties of NOS 1.1.1 Synthesis of NO NO is produced from L‐arginine (L‐Arg) by NOS. Three distinct isoforms of this enzyme have been described with different gene products, different localization, regulation, and calatalytic properties. The endothelial NOS (eNOS, NOS‐3, NOS‐III, type III) – first found in endothelial cells – is responsible for cardiovascular actions, the neuronal NOS (nNOS, NOS‐1, NOS‐I, type I) – first found in neurons – plays a role in the intercellular communication of neurons, and the inducible NOS (iNOS, NOS‐2, NOS‐II, type II) – first found in immune cells (e.g., macrophages) – which is mainly involved in cellular defense mechanisms (Alderton et al., 2001). Since their first discovery, it has been shown that all isoforms can be found in a wide variety of cells but the original names remained unchanged. Two isoforms, nNOS and eNOS are constitutive enzymes that are always present in the cells capable for their expression, while iNOS is inducible and appears only in response to an immunological stimulus. The NO synthesizing enzymes were described in 1989 and the isoforms were cloned between 1991 and 1994. These isoforms show 51–57% homology in human (nNOS 161 kD, eNOS 133 kD, iNOS 131 kD). Some parts of the enzyme isoforms show structural homology with cytochrome P450 reductase as well (> Figure 16-1). Structurally these enzymes have two subunits, a reductase and an oxygenase domain; however, for the synthesis of NO a dimerization is necessary, that is, in a functional complex there are two reductase and oxygenase domains. In addition, for the stabilization of this complex, two activated calmodulin molecules are necessary. In the case of nNOS and eNOS the association of calmodulins to the reductase domains occurs in response to the elevation of intracellular Ca2þ concentration and after the reduction of Ca2þ levels calmodulin dissociates

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. Figure 16-1 Structure of the human nitric oxide synthase (NOS) isoforms. All isoforms consist of a reductase and an oxygenase domain. The reductase domain shows high degree of amino acid sequence identity with the enzyme cytochrome P450 reductase. Binding site for the major cofactors (NADPH, FAD, FMN, THB and HAEM), calmodulin (CaM) and the substrate, L-arginine (L-Arg) are also shown. The neuronal NOS (nNOS) isoform contains a PDZ domain, which connects nNOS to the second PDZ domain of PSD-95. This protein connects nNOS to the intracellular region of NMDA receptors. In the inducible NOS (iNOS) there is a 40 amino acid deletion in the FMN binding domain. The lack of this autoinhibitory loop explains the Ca2+-independent operation of iNOS, because in the endothelial and neuronal isoforms the destabilizing effect of this loop results in the dissociation of calmodulin (and the concomitant inactivation of the enzymes) in the absence of Ca2+

from the complex resulting in an inactive conformation. The iNOS enzyme tightly and irreversibly binds calmodulin therefore the Ca2þ signal is not necessary for the activation; however, the dimerization of this isoform requires tetrahydrobiopterin (THB). The major function of all NOS isoforms is to catalyze a reaction in which NO, citrulline, and NADP is produced from L‐Arg, NADPH, and O2. During these reactions the enzymes utilize relatively tightly bound cofactors such as FAD, FMN, THB, and iron protporphyrin IX (haem). Electrons are provided by NADPH to the reductase domain of the enzyme and transferred to the oxygenase domain via the FAD and FMN redox chain. The reductase domain is very similar to cytochrom p450 reductase since it has the same electron transport function. FMN gives the electron to the haem group of the second enzyme in the dimer, the reason why the monomer form is not active. The electrons are interacting with the haem group and THB at the active site and catalyze the reaction of O2 with L‐Arg, generating citrulline and NO. The electron transport is possible only if the reductase and oxygenase domains are closely connected following the binding of Ca2þ/calmodulin complex. In addition, the oxygenase domain has a Zn2þ binding site. The role of Zn2þ is the stabilization of dimer structure. The catalytic site on the oxygenase domain of human iNOS and eNOS are very similar (that is why the development of a selective inhibitor is very difficult). The dimerization of iNOS is different

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from that of eNOS and nNOS, because in the case of iNOS only the oxygenase subunit is involved whereas the latter two isoforms associate through both the oxygenase and reductase subunits. That is why the activity of iNOS is dependent on THB (which cofactor is critical for the oxygenase subunit dimerization). The dimerization is a critical precondition of enzyme activity. The basic reaction catalyzed by the enzyme is the following: L-Arg

þ NADPH þ O2 ¼ citrulline þ NADP þ NO or NO
ðnitroxyl anionÞ

A clearcut 1:1 stoichiometry of NO and citrulline could not be shown so far, therefore the most accepted idea is that both NO and NO
is synthesized by NOS. It is an interesting observation that without superoxide dismutase (SOD) no authentic NO can be detected. Some data suggest that NO
is the first product which is oxidized by SOD resulting in NO. The other explanation that SOD removes superoxide from the environment, which prevents the formation of peroxynitrite (ONOO
). Under certain conditions NOS can synthesize superoxide because an uncoupled NADPH oxidation can result in the production of this compound. This kind of reaction is especially typical for nNOS, when the major substrate, L‐Arg or THB is not available in sufficient amount or in the presence of some NOS inhibitors. Simultaneous synthesis of NO and superoxide might lead to the instantaneous production of peroxynitrite.



NO þ superoxide O
2 ¼ peroxynitrite ðONOO Þ or NO þ O2 ¼ peroxynitrite ðONOO Þ This feature of NOS enzymes might explain the differences in the effect of endogenous NO synthesized by NOS isoforms and exogenous NO released by NO donors. The latter compounds always produce NO, while the synthesis might produce a number of different reactive nitrogen species (RNS). It has been shown that eNOS is less inclined to produce RNS other than NO whereas iNOS and nNOS have the propensity for this.

1.1.2 Regulation of NOS Activity NO has the potential to inhibit its own synthesis via an interaction with the haem group of the oxygenase domain. That is the mechanism by which NO can inhibit other haem containing enzymes as well. This autoinhibition is weak in iNOS which makes its continuous high activity possible. As we mentioned previously, the Ca2þ/calmodulin complex is necessary for the activation of eNOS and nNOS but not required for iNOS. The reason for this phenomenon is an autoinhibitory loop in nNOS and eNOS in the middle of the FMN segment, which destabilizes the calmodulin binding in the absence of Ca2þ. In the absence of calmodulin the electron transfer is not possible between FMN and haem, because in this conformation the distance of oxygenase and reductase subunits is too large. Another regulatory mechanism is the phosphorylation of nNOS and eNOS. The catalytic activity of these isoforms is decreased following phosphorylation by cAMP‐dependent protein kinase, protein kinase C, or Ca2þ/calmodulin‐dependent protein kinase II.

1.1.3 Subcellular Localization of NOS Acylation by myristate and palmytate is required for efficient localization of eNOS to the plasmalemmal caveolae of endothelial cells, where eNOS binds to caveolin‐1, whereas in cardiac myocytes to caveolin‐3. Endothelial NOS is inhibited by caveolin‐1 and this inhibition is regulated by Ca2þ/calmodulin. The N‐terminal of nNOS is unique, because it has a PDZ domain, which targets nNOS to the second PDZ domain of PSD‐95 protein in the brain. This organization is very important, because PSD‐95 can also bind to the NMDA receptors, which makes a very close connection between glutamatergic activation and NO production (see later). Splice variants of nNOS (b and g) lack this PDZ domain and occur in soluble form. Peptides that bind to either nNOS PDZ domains or PSD‐95 can uncouple nNOS activity from NMDA receptor stimulation. One of this regulatory protein is the protein inhibitor of nNOS (PIN), which can bind to the N‐terminal region of nNOS. Recently it has been proposed that PIN is not a specific inhibitor of

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nNOS but rather an axonal transport protein. Another regulatory protein is the carboxy‐terminal PDZ ligand of nNOS (CAPON), which binds to the PDZ domain and prevents the association of nNOS with the PSD‐95/NMDA complex.

1.1.4 Expression of NOS Splice Variants Posttranscriptional modification of nNOS has been described. Neuronal NOSa is the full length enzyme, nNOSb, nNOSg, nNOSm, and nNOS‐2 are the shorter splice variants. They were discovered in nNOS knockout mice, because following exon 2 deletion (PDZ domain encoding region) a residual NOS activity has been observed. Neuronal NOSb and nNOSg, described in nNOS knockout animals, do not bind to the PSD95/NMDA complex. Their function is unknown. The catalytic activity of nNOSb is 80%, but the nNOSg form is almost inactive (3% as compared to the nNOSa form). The nNOSm variant has an insertion, it is selectively expressed in rat heart and skeletal muscle, and in human and rat penis and urethra. Its activity is similar to that of nNOSa. Neuronal NOS‐2 is a shorter form but it has a deletion inside the sequence. The L‐Arg binding site is deleted, therefore most probably this variant is catalytically inactive; however its enzymatic activity has not been investigated so far. This form might have some regulatory function on the nNOSa form. Little is known about iNOS splice variants and eNOS splice variants have not been found so far.

1.1.5 NOS Inhibitors The inhibitors of NOS are widely used for studying the nitrergic mechanisms and the isoform selective compounds might have important therapeutic implications in the future. The NOS inhibitors interact with the enzyme on several locations. The competitive inhibitors of NOS associate to the L‐Arg binding site and usually they are L ‐Arg analogues like N(omega)‐nitro‐ L‐arginine methyl ester (L ‐NAME), N(omega)‐nitro‐ L‐arginine (L‐NNA), N(omega)‐monomethyl‐L‐arginine (L‐NMMA). Others have more complex mechanism of action because they bind to the L‐Arg site and undergo covalent changes like aminoguanidine or the acetamidine inhibitors such as (5)‐(1‐iminoethyl)‐ornithine (L‐NIO), L‐N6‐iminoethyl‐lysine (L‐NIL). Further target might be the THB binding site, where the pterine analogues like 7‐nitro indazole (7‐NI) can act, the haem group, where the haem‐binding inhibitors (such as imidazole) take effect, and the flavoprotein and calmodulin‐binding site. The specificity of these inhibitors is a key question if the role of specific NOS isoforms is the subject of investigation. Selectivity depends on a number of factors, such as kinetic properties (time‐dependence), way of administration, estimation of effects on other targets. For example, 7‐NI is used as a specific nNOS inhibitor; however, this is true only for in vivo administration, because in vitro this compound is not selective. The arginine analogues are not selective inhibitors but L‐NIO is believed to be eNOS selective, whereas L‐NIL is relatively specific for iNOS. A really selective iNOS inhibitor could be theoretically very useful for the treatment of septic shock, since in this case the overexpression and overactivation of iNOS leads to a fatal circulatory collapse because of the maximal relaxation of blood vessels, nevertheless, a clinically acceptable compound has not been developed so far.

1.2 Effector Mechanisms of NO A characteristic feature of the nitrergic systems is that a unique and exclusive mechanism does not exist for the mediation of the actions of NO. At molecular level there are two important reactions, which provide basis for NO to influence the function of a wide range of proteins: the binding to transition metal centers (like in the case of haem containing enzymes) and the reaction with cysteine thiol groups, the so‐called S‐nitrosylation (Ahern et al., 2002). The best known and first recognized mechanism, the activation of soluble guanylyl cyclase (sGC) belongs to the first reaction type, because sGC has a prosthetic haem group and NO binds to this site. Although sGC is sometimes referred as the NO receptor, this protein has a much

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wider function than simply to detect and react to NO with an increased production of the cyclic nucleotide cGMP. In the case of eNOS the elevation of cGMP (via more intermediate steps) leads to the relaxation of smooth muscles in the wall of blood vessels and the concomitant decrease of blood pressure, but in the nervous system many effects of NO produced by nNOS are mediated also via the sGC/cGMP pathway. Immunohystochemical studies have revealed that the distribution of sGC is complementary to nNOS, providing convincing evidence for the functional cooperation of these enzymes.

1.3 NO and nNOS in the Central Nervous System 1.3.1 General Features It has been shown that although nNOS is present also in glia and cerebral blood vessels, it is predominantly located in neurons. Although nNOS positive neurons represent only approximately 1% of nerve cells, they usually possess very rich axon arborization and the nNOS is expressed in sufficient amount in dendrites and axons, therefore almost all neurons in the brain are exposed to NO produced by nNOS containing nerve cells. NO has dramatically changed the classical concept of neurotransmission. Although there is a specific enzyme that synthesizes the molecule, NO cannot be stored in neurons but it is immediately released upon production and readily diffuses away from the site of synthesis. There is no specific NO receptor but NO can modify the function of a number of proteins via reversible chemical reactions (like S‐nitrosylation or association with prosthetic metal groups). Although all isoforms of NOS can be found in the central nervous system (CNS), because of the temporal and spatial properties (timing of activation, regional distribution) the specific actions on neurotransmission may be attributed primarily to NO produced by nNOS located in neurons. It has been observed that in the CNS the Ca2þ/calmodulin‐dependent nNOS produces NO almost exclusively following activation of NMDA receptors. Since a wide range of receptors (e.g., nicotinic acetylcholine receptors, 5‐HT3 receptors, etc.) may mediate effects resulting in increased intracellular Ca2þ concentration, the close relationship between NMDA receptor activation and NO synthesis seemed to be puzzling; however, clarification of the molecular organization of glutamatergic synapses provided explanation for this phenomenon. It has been shown that nNOS is connected to the NMDA receptors via a postsynaptic density protein (PSD‐95) thus the enzyme is directly exposed to the flux of Ca2þ ‐ions entering the ion channel of activated NMDA receptors. Ca2þ ‐transients arising due to the activation of other receptors presumably are too diluted by the time they reach the vicinity of the enzyme, therefore nNOS can be ‘‘switched on’’ only by NMDA receptors. Consequently, the level of endogenously produced NO around the NMDA receptor/ nNOS‐containing synapses reflects the activity of glutamatergic neurotransmission. This might be very useful from the point of view of interneuronal communication, since the concentration of NO is a reliable and interpretable signal for the surrounding neurons. The potential significance of this kind of interaction can be demonstrated by two examples, the role of NO in the long‐term potentiation (LTP) and in the nonsynaptic communication between glutamatergic and monoaminergic neurons.

1.3.2 NO and LTP It is generally accepted that the cellular basis of learning and memory formation is the activity dependent and permanent increase in the synaptic efficacy between neurons. This kind of synaptic plasticity is called LTP and it has been observed in glutamatergic synapses. The intensive research of this phenomenon revealed that the increased efficacy at a given synapse consists of both presynaptic and postsynaptic events. In addition, it was clear that the primary change in the postsysnaptic element has to be somehow detected by the presynaptic site in order to develop the necessary changes, that is, there must be some information flow in the opposite direction within the synapse. Soon after the discovery of the close connection between the NO synthesis and NMDA receptor activation, it has been proposed that NO might act as a retrograde

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messenger, which influences synaptic transmission in the presynaptic cell and promotes synaptic plasticity (Holscher et al., 1997). Accumulating evidence supports this hypothesis because the inhibition of NO synthesis or the genetic deletion of nNOS prevents or significantly decreases the development of LTP. In addition, it has been clarified that the presynaptic action of NO is mediated through the sGC/cGMP pathway, because the inhibitors of sGC also reduced LTP. Nevertheless, the picture is more complicated because NO‐independent forms of LTP have also been described. It has been suggested that not only nNOS but also eNOS is involved in the process because a complete inhibition of LTP could be achieved in certain brain areas only in double knockout mice lacking both nNOS and eNOS. The most probable explanation is that there are alternative mechanisms for the induction of LTP, which might be regionally different in the brain and some of them are NO‐dependent whereas others are not. Consistent with these data, the manipulation of endogenous NO production affects some types of learning in animal experiments.

1.3.3 NO as a Nonsynaptic Link between Glutamatergic and Monoaminergic Neurons Glutamate is the major excitatory transmitter of the brain. In contrast to other transmitters, glutamate participates mainly in synaptic interactions, because glutamatergic release sites are located predominantly within synapses. Although some synaptic spillover of glutamate has been shown at room temperature, this spillover almost completely disappeared at 37  C due to the very efficient neuronal and glial uptake mechanisms. However, even if spillover exists at body temperature, the diffusion of escaped glutamate is very limited because of the neuronal and glial uptake processes, therefore under physiological conditions only a small portion of released glutamate actually leaves the synaptic cleft and may reach only glutamate receptors located perisynaptically or in neighboring synapses. However– due to its physicochemical properties – NO is an ideal mediator of nonsynaptic interactions (Vizi, 1984). Since the lipid membranes cannot form a barrier for NO, in spite of its very short (few seconds) half‐life it can diffuse away a few hundred from the site of production. Comparing this distance to the width of a synaptic cleft (20 nm) or the size of a cell body (few mm) it is evident that NO produced at a glutamatergic synapse may influence the function of a very large number of nerve cells in a sphere around the activated synapse. It has been observed that NO is able to inhibit the function of monoamine transporters. The inhibitory action of NO could be revealed not only by NO donors in synaptosomal preparations but also by NOS inhibitors in functional in vitro and in vivo release experiments, indicating that endogenous NO exerts a tonic inhibitory effect on transporters. The molecular mechanism of inhibition is the S‐nitrosylation of a cysteine thiol group in the seventh transmembrane domain of monoamine transporters (Kaye et al., 2000). The effect of NO on monoamine uptake allows NO to signal glutamatergic activity to the environment through the change of inhibitory tone on transporters (> Figure 16-2). This effect may increase the concentration of monoamines in the extracellular space by prolonging their lifetime in a local volume around the synapse, and this rise represents the specific response of monoaminergic systems to the activation of glutamatergic neurotransmission. Since monoamines can inhibit the release of glutamate, this NO‐mediated interaction may serve as a negative feedback loop. Thus, with the help of NO, glutamate may participate in long‐range nonsynaptic interactions. A very peculiar feature of the NO‐mediated communication is that the information originating from the glutamatergic system can reach the monoaminergic system even if the recipient cells, in contrast to the previously known synaptic and nonsynaptic interactions, do not express specific receptors sensitive to glutamate. The monoaminergic neurons have to express only their ‘‘own’’ proteins, the transporters, which enables them to receive a message from another neurotransmitter system. Since the monoaminergic systems can respond to the activation of glutamatergic transmission without receiving synaptic input and without expressing glutamate receptors, the effect of NO on transporters represents a new form of interneuronal communication, a nonsynaptic interaction without receptors (Kiss and Vizi, 2001).

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. Figure 16-2 NO-mediated nonsynaptic communication between glutamatergic and monoaminergic systems. Release of glutamate stimulates NMDA receptors and the concomitant influx of Ca2+-ions (not shown) activates nNOS coupled to the receptor via PSD-95. NO synthetized by the enzyme spreads over in a sphere and reaches monoaminergic vaicosities in the environment of activated synapses. The actual extracellular concentration of monoamines depends on the balance of release and uptake processes. The nonsynaptic signal, i.e., the appearance of NO inhibits the function of transporters (T), which increases the extracellular concentration of monoamines in a local volume around the activated glutamatergic synapse even if the amount of released monoamines is unchanged. The elevated monoamine level exerts an inhibitory effect on glutamate release via presynaptic monoamine receptors (MAR) located on glutamatergic terminals. Modified with permission from Kiss and Vizi, 2001

1.4 NO and Neurotoxicity NO is a very reactive free radical that can be harmful for cells. Exactly this feature is used in the immune response, when macrophages destroy the pathogenic organisms and defected cells by the overproduction of NO synthesized by iNOS. In the CNS, NO might be a useful physiological messenger or a dangerous pathological agent depending on the actual cellular environment, and the rate and amount of NO production. At lower concentration (below 1 mM), typically occurring in unstressed cells, NO exist at a redox state that does not readily form damaging RNS. Under these conditions the superoxid concentration is low, therefore NO cannot form peroxynitrite with this compound. In contrast, neuroprotective processes are activated by NO (e.g., BDNF is upregulated, antiapoptotic genes are activated, and caspase activation is inhibited). Usually low dose of NO is released by the Ca2þ ‐dependent constitutive NOS enzymes (eNOS, nNOS). Cells have certain natural resistance mechanisms against the nitrooxidative stress exerted by free radicals. The protective mechanisms involve the activation of SOD enzyme, which converts superoxide anions into H2O2, glutathione/glutathionyl peroxidase activation that reduces superoxide levels, thioredoxin reductase/thioredoxin system activation, upregulated proteolysis of damaged proteins, increased DNA repair activity, and hemoxygenase upregulation. The physiological levels of NO do not exceed the protective capacity of these mechanisms. What is more, low concentration of NO might induce changes in

Nitric oxide and other diffusible messengers

16

the cells which make them more resistant to subsequent toxic concentrations of NO. This phenomenon is the induced adaptive resistance, which can be a useful neuroprotective mechanism. NO‐induced neurotoxicity occurs when the concentration of NO exceeds the normal level (about 1 mM). Under these conditions the generated NO and associated RNS are harmful for the neurons. The abnormal amount of NO might be produced in CNS injury and inflammation by macrophages that infiltrate the lesioned area and by activated microglia and astrocytes. These cells express iNOS in response to injury or inflammation and produce 2–3 orders of magnitude higher amount of NO. During ischemic episodes neurons might also produce extreme amount of NO because of the high extracellular concentration of excitatory amino acids, primarily glutamate, which overactivates NMDA receptors and the associated nNOS. In neurotoxicity models nNOS induces cell damage while eNOS is rather neuroprotective because of the amelioration of cerebral blood flow by the cGMP‐mediated relaxation of blood vessels. It has been observed that nNOS knockout mice are less sensitive for cerebral ischemia, they show less brain damage, while eNOS knockout mice are more vulnerable. These opposite functions provide explanation for the often contradictory results on the neuroprotective or neurotoxic role of NO in the CNS. Oxydative cell damage can be seen not only in CNS injury, inflammation or ischemia but also in neurodegenerative states like Alzheimer’s disease or Parkinson’s disease, where the trace of nitrated proteins can be detected. Multiple Sclerosis and ALS are also coupled to nitrooxidative stress. Under these conditions the abnormal level of nitrooxidative species cannot be handled by the natural protective mechanisms of neurons. The mechanisms of cell damage induced by NO and RNS are complex. The lipid peroxidation destroys biological membranes, the free radicals (primarily peroxynitrite) induce DNA damage, which activates the nuclear repair enzyme polyADP‐ribose synthase (PARS). Overactivation of PARS depletes energy stores, which leads to cell death. Inhibition of mitochondrial respiratory chain because of the irreversible binding of NO to mitochondrial cytochromes exacerbates the depletion of energy stores. Enzyme nitrosylation (e.g., phosphokinase C, glyceraldehydes‐3‐phosphate dehydrogenase) impairs cell viability. Resistance genes (e.g., BDNF expression) are inhibited by high concentration of NO. The increased intracellular haem release results in the appearance of redox‐active iron, which leads to the concomitant formation of further free radicals (e.g., hydroxyl groups). In the presence of available iron and superoxide, NO forms peroxynitrite anion, which reacts with nitrate tyrosine residues forming 3‐nitrotyrosine, thereby destroying structure and function of tyrosine‐rich proteins, like structural neurofilaments resulting in impaired axonal trafficking. The DNA damage also induces apoptosis because of the activation of caspase system.

1.5 NO in the Peripheral Nervous System It is now well‐accepted that NO has an important role in the periphery as a neurotransmitter in non‐ adrenerg non‐cholinerg (NANC) nerves (Esplugues, 2002). The main difference between the operation of central and peripheral nNOS is that the former is activated by Ca2þ ‐influx through the NMDA receptors, whereas the latter is turned on because of the opening of voltage‐sensitive calcium channels since it is not associated with NMDA receptors. The major function of nitrergic nerves is to induce relaxation of smooth muscle in the gastrointestinal, respiratory, and urogenital tract. These nerves release NO or closely related redox product from NO. There is some controversy as to whether NO is the primary transmitter or NO production is necessary for the release of VIP, which would be the final mediator of smooth muscle contraction. In the gastrointestinal system smooth muscle cells contain sGC in the vicinity of nNOS positive neurons. The oesophageal and pyloric sphincters are operated via NANC nerves. The myenteric plexus also contains nNOS positive cells. The lack of these cells results in delay of colonic transit. In the pulmonary tract nitrergic nerves are believed to be the major nervous bronchodilatator pathway in humans. NO inhalation proved to be useful in acute respiratory distress syndrome, and other restrictive pulmonary diseases. In the vascular system nNOS is found in the perivascular nerves and provides an alternative mechanism for the blood vessel relaxation in addition to the primary eNOS regulation. In the urogenital tract there is a role for spinal cord NO in micturition reflex. Nitrergic mechanism plays an important role also in the penile erection because the relaxation of corpus cavernosum is mediated via NANC nerves. The cGMP signal is terminated in the corpus cavernosum by a specific isoenzyme, phosphodiesterase‐5.

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Selective inhibition of this enzyme by sildenafil (Viagra) or zaprinast restores erectile function because of the prolongation of the cGMP signal, which extends the relaxation of corpus cavernosum. NANC nerves also innervate clitoral corpus cavernosum. In the skeletal muscle high levels of nNOS is expressed by muscle cells, especially the muscle‐specific splice variant nNOSm. Here NO might be the regulator of contractile force.

2

Carbon Monoxide

The appearance of NO as a possible neuromodulator initiated an intensive search for other diffusible molecules that could play similar role in the nervous system. Because of the significant chemical similarity carbon monoxide (CO) was among the candidates. Accumulating data indicate that CO is able to fulfil very similar functions in the CNS as NO. CO is produced by heme oxygenase (HO), which cleaves the haem ring into CO and biliverdin. The latter compound is rapidly reduced to bilirubin by biliverdin reductase. There are two isoforms of HO. The most abundant form is HO1, which is found in the periphery, mainly in the spleen and degrades senescent red blood cells. HO1 is an inducible enzyme, its expression is increased by haem or oxidative stress. The other isoform, HO2 is constitutive and selectively localized to the brain. In situ hybridization studies revealed that the localization of HO2 more closely resembles those of sGC than the localization of nNOS. This is understandable because the action of CO is very similar to that of NO, it binds to the haem group of sGC and increases the production of cGMP. The neurotransmitter role of CO has been established first in the myenteric plexus, where the localization of nNOS and HO2 is essentially identical, and the NANC relaxation of the intestine has both a CO‐ and a NO‐mediated component, as it was revealed by pharmacological agents and also in genetically modified (HO2
/
, nNOS
/
and double knockout) animals (Snyder et al., 1998). Nevertheless, there are important differences between nNOS and HO2, because the former is activated by the rise of Ca2þ ‐concentration in the vicinity of the enzyme, whereas the latter is activated by protein kinase C, which phosphorylates HO2. Since HO2 is especially concentrated in hippocampal pyramidal cells, it has been suggested that CO, similarly to NO, might be a retrograde messenger in LTP (Hawkins et al., 1994). Experimental data support this hypothesis, because the HO inhibitor zinc protoporphyrin IX blocked the induction of LTP in hippocampal slices, and long‐lasting increase in the amplitude of evoked potentials were observed when CO was applied at the same time as weak tetanic stimulation (Zhuo et al., 1993).

3

Hydrogen Peroxide

Hydrogen peroxide (H2O2) is a metabolic byproduct of the reactive oxygen species (ROS) elimination, since it is produced by (SOD) from superoxide anion (O
2 ), which originates from the incomplete reduction of O2, and might be synthesized also by NOS, especially by nNOS under special conditions (see above). In the final step of ROS elimination H2O2 is converted to harmless endproducts, H2O and O2 by catalase and gluthation peroxidase. H2O2 is a membrane‐permeable molecule and theoretically it could be a diffusible messenger like NO or CO. Recently it has been shown that glutamate, acting at AMPA receptors, inhibits DA release in the striatum by a mechanism mediated by H2O2. The inhibition on DA release is exerted by the activation of presynaptic ATP‐sensitive potassium channels, which are opened by H2O2. This kind of interaction provides a novel mechanism for the intercellular communication between neurons by means of a diffusible messenger, H2O2 (Avshalumov et al., 2003).

References Ahern GP, Klyachko VA, Jackson MB. 2002. cGMP and S‐nitrosylation: Two routes for modulation of neuronal excitability by NO. Trends Neurosci 25: 510-517.

Alderton WK, Cooper CE, Knowles RG. 2001. Nitric oxide synthases: Structure, function and inhibition. Biochem J 357: 593-615.

Nitric oxide and other diffusible messengers Avshalumov MV, Chen BT, Marshall SP, Pena DM, Rice ME. 2003. Glutamate‐dependent inhibition of dopamine release in striatum is mediated by a new diffusible messenger, H2O2. J Neurosci 23: 2744-2750. Esplugues JV. 2002. NO as a signalling molecule in the nervous system. Br J Pharmacol 135: 1079-1095. Hawkins RD, Zhuo M, Arancio O. 1994. Nitric oxide and carbon monoxide as possible retrograde messengers in hippocampal long‐term potentiation. J Neurobiol 25: 652-665. Holscher C. 1997. Nitric oxide, the enigmatic neuronal messenger: Its role in synaptic plasticity. Trends Neurosci 20: 298-303. Kaye DM, Gruskin S, Smith AI, Esler MD. 2000. Nitric oxide mediated modulation of norepinephrine transport: Identification of a potential target for S‐nitrosylation. Br J Pharmacol 130: 1060-1064.

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Kiss JP, Vizi ES. 2001. Nitric oxide: A novel link between synaptic and nonsynaptic transmission. Trends Neurosci 24: 211-215. Palmer RM, Ferrige AG, Moncada S. 1987. Nitric oxide release accounts for the biological activity of endothelium‐derived relaxing factor. Nature 327: 524-526. Snyder SH, Jaffrey SR, Zakhary R. 1998. Nitric oxide and carbon monoxide: Parallel roles as neural messengers. Brain Res Rev 26: 167-175. Vizi ES. 1984. Non‐synaptic interaction between neurons: Modulation of neurochemical transmission. New York: Wiley. Zhuo M, Small SA, Kandel ER, Hawkins RD. 1993. Nitric oxide and carbon monoxide produce activity‐dependent long‐term synaptic enhancement in hippocampus. Science 260: 1946-1950.

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Molecular Organization and Regulation of Glutamate Receptors in Developing and Adult Mammalian Central Nervous Systems

E. Molna´r

1

Historical Overview . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 416

2 2.1 2.2

Molecular Organization and Heterogeneity of Glutamate Receptors . . . . . . . . . . . . . . . . . . . . . . . . . 417 Ionotropic Glutamate Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 417 Metabotropic Glutamate Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 420

3 3.1 3.2 3.3 3.4

Developmental Changes in Glutamate Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 421 Developmental Changes in AMPA Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 421 Developmental Changes in NMDA Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 422 Developmental Changes in Kainate Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 422 Developmental Changes in Metabotropic Glutamate Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 423

4 4.1 4.2

Distribution of Glutamate Receptors in the CNS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 423 Synaptic Distribution of Glutamate Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 423 Extrasynaptic Glutamate Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 426

5 5.1 5.1.1 5.1.2 5.1.3 5.1.4 5.2 5.2.1 5.2.2 5.2.3 5.2.4

Regulation of Glutamate Receptors during Synaptic Plasticity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 427 Phosphorylation of Glutamate Receptors and Synaptic Plasticity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 427 Phosphorylation of AMPA Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 427 Phosphorylation of Kainate Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 429 Phosphorylation of NMDA Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 429 Phosphorylation of mGluRs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 429 Receptor Trafficking and Synaptic Plasticity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 430 Recruitment of AMPA Receptors at Synapses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 430 Kainate Receptor Trafficking . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 432 NMDA Receptor Trafficking . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 433 Trafficking of mGluRs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 434

6

Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 434

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Abstract: The amino acid glutamate is the major excitatory neurotransmitter in the mammalian central nervous system (CNS), and it exerts its physiological effects by binding to a number of different ionotropic (ligand‐gated ion channels) and metabotropic (G‐protein‐coupled) glutamate receptors. In addition to excitatory neurotransmission, glutamate receptors play an essential role in neuronal differentiation, plastic changes in efficacy of synaptic transmission, neurodegeneration, and neuronal cell death. The application of molecular cloning technology identified a complex and diverse receptor family. To date, 18 different ionotropic glutamate receptor (iGluR) genes and 8 genes for metabotropic glutamate receptors (mGluRs) have been identified from mammals. The availability of more selective drugs, gene knockout mice and high‐ resolution immunohistochemical studies started to reveal the pharmacological and functional properties of individual iGluR subunits and mGluR isoforms. Wide range of studies in recent years indicated that the activity and synaptic distribution of various glutamate receptors are dynamically regulated by phosphorylation and protein–protein interactions, which are key mechanisms in mediating synaptic plasticity. This chapter reviews some of the recent progress in glutamate receptor research with special emphasis on the molecular diversity of this receptor system and its implications for neuronal development and synaptic plasticity. List of Abbreviations: ABP, AMPAR‐binding protein; AMPA, a‐amino‐3‐hydroxy‐5‐methyl‐4‐isoxazole propionate; CaMKII, Ca2þ/calmodulin‐dependent protein kinase II; CNS, central nervous system; GABA, g‐aminobutyric acid; GRIP1, glutamate receptor interacting protein 1; iGluR, ionotropic glutamate receptor; IP3, inositol 1,4,5‐trisphosphate; LTD, long‐term depression; LTP, long‐term potentiation; mGluR, metabotropic glutamate receptor; MLCK, myosin light‐chain kinase; NMDA, N‐methyl‐D‐aspartate; NSF, N‐ethylmaleimide‐sensitive factor; PI3K, phosphatidylinositol 3‐kinase; PICK1, protein interacting with C‐kinase 1; PKA, protein kinase A; PKC, protein kinase C; PLC, phospholipase C; PSD, postsynaptic density; RT‐PCR, reverse transcriptase polymerase chain reaction; SAP, synapse‐associated protein

1

Historical Overview

The realization in the 1930s that there is a relatively high concentration of glutamate in the brain prompted early speculations of an important neurophysiological role of the amino acid, and this in turn led to a variety of trials in the 1940s of dietary glutamate and glutamine in the treatment of learning disorders and epilepsy. The role of glutamate in the brain, aside from the obvious one as a protein constituent, was at the time considered more in terms of energy metabolism, given the close association of the amino acid with the citric acid cycle (Watkins and Jane, 2006). Glutamatergic synaptic transmission in the mammalian central nervous system (CNS) was slowly established over a period of some 20 years, dating from the 1950s. An early indication of a special role of glutamate in electrophysiological processes was the observation that injection of glutamate into brain or carotid arteries produced convulsions (Hayashi, 1954), which led to the speculation that glutamate was a transmitter in the mammalian CNS. In the late 1950s, initial studies indicated that L‐glutamate and a number of other naturally occurring acidic amino acids excited single neurons in the mammalian brain (Curtis et al., 1959). It is now commonly believed that L‐glutamate is the principal mediator of fast excitatory neurotransmission among vertebrate neurons. The emergence of increasingly specific pharmacological tools during the 1970s started to reveal considerable functional diversity. The family of glutamate‐activated cation channels (ionotropic glutamate receptors [iGluRs]) was classified into three major pharmacological subfamilies, defined by their most selective agonists: a‐amino‐3‐hydroxy‐5‐methyl‐4‐isoxazole propionate (AMPA), kainate, and N‐methyl‐D‐ aspartate (NMDA) receptors. In the mid‐1980s, evidence began to appear for the existence of another glutamate receptor group termed metabotropic glutamate receptors (mGluRs) that are directly coupled to second‐messenger systems via GTP‐binding proteins (Sladeczek et al., 1985; Nicoletti et al., 1986). However, glutamate receptor proteins remained elusive until the late 1980s. This was due to the lack of sufficiently specific and high‐affinity receptor ligands and the low abundance of glutamate receptors in the brain, which hindered biochemical isolation efforts and prevented the use of conventional (partial sequence‐ based) cloning strategies. The application of the newly emerging expression cloning approach lead to a

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breakthrough and provided the original sequence information for the first iGluR subunits GluR1 (Hollmann et al., 1989) and NR1 (Moriyoshi et al., 1991). The first member of the family of mGluRs (mGluR1a) was also discovered by expression cloning (Houamed et al., 1991; Masu et al., 1991). These sequence information in combination with other cloning techniques, such as low‐stringency hybridization screening and reverse transcriptase polymerase chain reaction (RT‐PCR) with degenerate primers, lead to the rapid identification of related iGluR subunits, mGluR isoforms and their splice variants (reviewed in Hollmann and Heinemann, 1994; Conn and Pinn, 1997). The precise identification of discrete iGluRs and mGluRs, the receptor localization studies, and the development of a range of transgenic animals lead to major advances in our understanding of glutamatergic signaling mechanisms in the CNS. The study of protein–protein interactions, phosphorylation– dephosphorylation reactions, and receptor trafficking is a rapidly developing area of growing importance with respect to glutamate receptor function (Collingridge et al., 2004). These phenomena are considered particularly relevant in the synaptic plasticity field.

2

Molecular Organization and Heterogeneity of Glutamate Receptors

2.1 Ionotropic Glutamate Receptors Pharmacological, biophysical, and molecular data support the existence of three main iGluR families, named after the agonists that preferentially stimulate them: AMPA, kainate, and NMDA. While there is a clear pharmacological boundary between NMDA receptors (NMDARs) and other iGluRs, AMPA receptors (AMPARs) and kainate receptors (KARs) share several agonists and antagonists, which can interact with both receptors (Dingledine et al., 1999). The development of new drugs with higher subtype and subunit specificity (> Figure 17‐1) and the availability of mice that are deficient for individual iGluR subunits have been fundamental to our progress in understanding the role of iGluRs in synaptic physiology. For example, the development of AMPAR antagonists 2,3‐benzodiazepines (e.g. GYKI 52466 and GYKI 53655) allowed the selective investigation of native AMPARs and KARs in neurons (Vizi et al., 1996; Tarnawa and Vizi, 1998; Lerma, 2003). The cloning of many iGluR subunits (> Figure 17‐1) has validated the initial pharmacological classification (Hollmann, 1999). AMPARs are built from homomeric or heteromeric combinations of subunits GluR1–4. Cloned KAR subunits are subdivided into low‐affinity (GluR5–7) and high‐affinity (KA‐1 and KA‐2) subunits. Electrophysiological and biochemical analysis of recombinant KARs indicates that like AMPARs, functional KAR channels are formed by both homomeric and heteromeric expression of GluR5–7 subunits. In contrast, the KA‐1 and KA‐2 subunits do not form functional homomeric channels, but they coassemble with the GluR5–7 subunits (Gallyas et al., 2003; Lerma, 2003). NMDARs are formed from hetero‐oligomeric assemblies of NR1 subunits with NR2 (A–D) and NR3A (Monyer et al., 1994). In addition, NR3A can assemble with NR1 (without other NR2 subunits) to form functional glycine receptors (Sasaki et al., 2002). Eight possible variants of the NR1 subunit arise by alternative splicing of a single gene transcript. The insertion of one splice cassette at the N‐terminal region of NR1 and the deletion of two independent consecutive splice variants at the C terminus have been described (Dingledine et al., 1999). There is general consensus that all iGluRs share a conserved transmembrane topology and stoichiometry. iGluRs are formed from the tetrameric assembly of homologous subunits around a central ion pore (Dingledine et al., 1999). The membrane topology of the iGluR subunits consists of a large extracellular N‐terminal domain and four hydrophobic membrane‐associated domains (M1–4; > Figure 17‐2). Early immunocytochemical and biochemical studies have indicated that unlike its location in other ligand‐gated ion channels, the C terminus is intracellularly located (Molna´r et al., 1993, 1994). Detailed analysis of membrane topology identified that iGluRs have only three transmembrane domains corresponding to M1, M3, and M4. M2 is a re‐entrant loop in the phospholipid bilayer, which does not span the membrane (Hollmann et al., 1994). The structure of M2 is similar to voltage‐gated Kþ channels and represents the channel pore‐forming region (M2, > Figure 17‐2). Each monomer carries its own ligand‐binding site, which consists of residues that are distributed throughout both the distal N‐terminal domain (called S1)

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and the extracellular loop between M3 and M4 (called S2) (> Figure 17‐2; Dingledine et al., 1999). The S1S2 ligand‐binding domain region of some of the iGluR subunits (GluR2, GluR5, GluR6, NR1, and NR2A) has been isolated and crystallized with and without bound ligand (Mayer and Armstrong, 2004; Mayer, 2005). X‐ray analysis of these proteins and protein–ligand complexes elucidated fine molecular details of the binding site, and the conformational changes of different agonists and antagonists induce to open or block the ion channels or modulate the manifestation of their activity (Mayer, 2006). Single‐particle electron microscopy provided some information about the conformational states of native AMPAR complexes and suggested that receptor desensitization is related to separation of the N‐terminal extracellular domains (Nakagawa et al., 2005). AMPARs are activated directly by glutamate binding, and their activation leads to changes in the membrane potential of the postsynaptic neuron. AMPARs are primarily permeable to Naþ, Kþ, and Ca2þ ions unless the edited form of GluR2 is present; in the later case, AMPARs are rendered impermeable to Ca2þ ions (Dingledine et al., 1999; Burnashev and Rozov, 2000). All AMPAR subunits undergo alternative splicing at the extracellular loop region (S2) adjacent to the last transmembrane domain (M4) yielding two splice variants termed flip and flop. The flip splice variants of GluR1–4 desensitize more slowly and to a lesser extent than the flop variants, which can influence the amplitude of the total AMPAR current. The C‐terminal tails of AMPAR subunits also undergo alternative splicing to yield short and long forms of the intracellular domain (Hollmann, 1999; Burnashev and Rozov, 2000). In AMPARs, GluR2 undergoes editing in the M2 channel pore‐forming region at the functionally significant Q/R site. Editing is a posttranslational

. Figure 17‐1 Classification of iGluR and mGluR families. In the dendrograms (left column), the length of the horizontal branches connecting any two subunits represents the degree of their relatedness based on pairwise sequence comparison. The scale bar at the bottom can be used to estimate the percentage sequence identity at the nodes that connect certain subsets of sequences. The known splice variants are indicated next to the code name of the iGluR subunits or mGluR isoforms. Key to selective ligands (left column): AMPA, (S)‐a‐amino‐3‐hydroxy‐5‐ methylisoxazole‐4‐propionic acid; LY404187, N‐2‐(4‐(4‐cyanophenyl)phenyl)propyl 2‐propanesulfonamide; (S)‐5‐fluorowillardiine, (S)‐(
)‐a‐amino‐5‐fluoro‐3,4‐dihydro‐2,4‐dioxo‐1(2H)pyridine propanoic acid; NBQX, 2,3‐dihydroxy‐6‐nitro‐7‐sulfamoyl‐benzo[f]quinoxaline; LY293558, (3S,4aR,6R,8aR)‐6‐[2‐(1(2)H‐tetrazole‐5yl) ethyl]decahydroisoquinoline‐3‐carboxylic acid; (S)‐ATPO, (S)‐2‐amino‐3‐[5‐tert‐butyl‐3‐(phosphonomethoxy)‐ 4‐isoxazolyl]propionate; ATPA, (RS)‐2‐amino‐3‐(3‐hydroxy‐5‐tert‐butylisoxazol‐4‐yl)propanoic acid; (S)‐5‐ iodo willardiine, (S)‐1‐(2‐amino‐2‐carboxyethyl)‐5‐iodopyrimidine‐2,4‐dione; LY339434, (2S,4R,6E)‐2‐amino‐4‐ carboxy‐7‐(2‐naphthyl)hept‐6‐enoic acid; SYM 2081, (2S,4R)‐4‐methylglutamic acid; LY382884, (3S,4aR,6S,8aR)‐ 6‐(4‐carboxyphenyl)methyl‐1,2,3,4,4a,5,6,7,8,8a‐decahydro isoquinoline‐3‐carboxylic acid; UBP302, (S)‐1‐ (2‐amino‐2‐carboxyethyl)‐3‐(2‐carboxybenzyl)pyrimidine‐2,4‐dione; NS3763, 4,6‐bis(benzoylamino)‐1,3‐benzenedicarboxylic acid; NMDA, N‐methyl‐D‐aspartic acid; tetrazolylglycine, (RS)‐(tetrazol‐5‐yl)glycine; (R)‐CPP, 3‐((R)‐2‐carboxypiperazin‐4‐yl)‐propyl‐1‐phosphonic acid; (1RS,10 S)‐PEAQX, (1RS,10 S)‐5‐phosphonomethyl‐1,4‐ dihydroquinoxaline‐2,3‐dione; PPDA, cis‐1‐(phenanthrene‐2‐carbonyl)piperazine‐2,3‐dicarboxylic acid; Ro 25‐6981, (aR,bS)‐a‐(4‐hydroxyphenyl)‐b‐methyl‐4‐(phenylmethyl)‐1‐piperidinepropanol maleate; L‐689,560, trans‐2‐carboxy‐5,7‐dichloro‐4‐phenylaminocarbonylamino‐1,2,3,4‐tetrahydroquinoline; MK‐801, dizocilpine/ (5S,10R)‐(þ)‐5‐methyl‐10,11‐dihydro‐5H‐dibenzo[a,d]cyclohepten‐5,10‐imine maleate; (S)‐3,5‐DHPG, (S)‐3, 5‐dihydroxyphenylglycine; CHPG, (RS)‐2‐chloro‐5‐hydroxyphenylglycine; DFB, 3,30 ‐difluorobenzaldazine/[(3‐ fluorophenyl)methylene]hyrazone‐3‐fluorobenzaldehyde; LY367385, (S)‐(þ)‐a‐amino‐4‐carboxy‐2‐methylbenzeneacetic acid; MCPG, (S)‐a‐methyl‐4‐carboxyphenyl‐glycine; MPEP, 2‐methyl‐6‐(phenylethynyl)pyridine hydrochloride; DCG IV, (2S,20 R,30 R)‐2‐(20 ,30 ‐dicarboxycyclopropyl)glycine; LY354740, (1S,2S,5R,6S)‐2‐aminobicyclo [3.1.0] hexane‐2,6‐dicarboxylate monohydrate; LY487379, N‐(4‐(2‐methoxyphenoxy)phenyl)‐N‐(2,2,2‐trifluoroethylsulfonyl)pyrid‐3‐ylmethylamine; LY341495, (2S)‐2‐amino‐2‐[(1S,2S)‐2‐carboxycycloprop‐1‐yl]‐3‐(xanth‐9‐yl) propanoic acid; EGlu, (S)‐a‐ethylglutamic acid; L‐AP4, (S)‐2‐amino‐4‐phosphonobutanoic acid; (S)‐3,4‐DCPG, (S)‐3,4‐dicarboxy‐phenylglycine; PHCCC, N‐phenyl‐7‐(hydroxyimino)cyclopropa[b]chromen‐1a‐carboxamide; CPPG, (RS)‐a‐cyclopropyl‐4‐phosphonophenylglycine. UBP1112, (RS)‐s‐methyl‐3‐methyl‐4‐phosphonophenylglycine. (a) Positive allosteric modulator, (b) negative allosteric modulator, and (c) NMDAR glycine site antagonist

Molecular organization and regulation of glutamate receptors in developing and adult mammalian CNS . Figure 17‐1 (Continued)

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. Figure 17‐2 Schematic representation of iGluR and mGluR membrane topology. See > Sections 2.1 and 2.2 for details. The extracellular N‐terminal domains of iGluRs and mGluRs contain glycosylation sites (Y)

change of one or more bases in the pre‐mRNA such that the codon(s) encoded by the gene and the codon(s) present in the mRNA differ. The arginin (R) codon (CGG) is not found in the GluR2 gene. It is introduced into the GluR2 mRNA by an adenosine to inosine conversion in the respective glutamine (Q) codon (CAG) of the GluR2 transcript by a double‐stranded RNA adenosine deaminase (Burnashev and Rozov, 2000). The inosine is subsequently read as a guanosine, resulting in the change of codon identity (CGG). The GluR2 subunit in the edited form is responsible for the Ca2þ impermeability of AMPARs (Hollmann, 1999; Burnashev and Rozov, 2000). At the postsynaptic level, KARs carry part of the synaptic charge. Presynaptically, KARs modulate transmitter release both at excitatory and inhibitory synapses. The diversity of KARs is increased by the existence of splice variants for GluR5 (1a–d, 2a–c), GluR6 (a–c), and GluR7 (a, b) receptor subunits (Lerma, 2003). Like the GluR2 AMPAR subunit, GluR5 and GluR6 are subjected to mRNA editing at the Q/R site. GluR6 can also undergo further editing at two more sites situated in the first hydrophobic M1 domain (Lerma, 2003). NMDARs require the binding of glutamate and the coagonist glycine, as well as membrane depolarization, to become activated and conduct primarily Ca2þ ions. The voltage‐dependent blockade of the NMDAR pore by Mg2þ produces this additional requirement for depolarization (Mayer et al., 1984; Nowak et al., 1984). While AMPARs are involved in the moment‐to‐moment information transfer between neurons, the major roles of NMDARs is to act as detectors of specific patterns of activity that trigger long‐ term changes in synaptic strength by modulating AMPAR responses. Activation of NMDARs can be achieved by sustained activation of AMPARs at the same synapse, resulting in membrane depolarization and relief of the voltage‐dependent Mg2þ block.

2.2 Metabotropic Glutamate Receptors The mGluRs are G‐protein‐coupled receptors, which mediate relatively slow responses to glutamate (Conn and Pinn, 1997; De Blasi et al., 2001). There are eight mGluR genes (mGluR1–8) and these have been divided into three groups based on pharmacological, signal transduction and sequence similarities (> Figure 17‐1). Group I comprises mGluR1 and mGluR5 and their splice variants (mGluR1a–d and

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mGluR5a,b), which are involved in the mobilization of intracellular Ca2þ by stimulating phospholipase C (PLC) and are activated by DHPG ((S)‐3,4‐dihydroxyphenylglycine) (Hermans and Challiss, 2001). In group II, the mGluR2 and mGluR3 are typically linked to inhibition of adenylyl cyclase (AC) and react effectively with DCG‐IV ((2S,10 S,20 S)‐2‐(carboxycyclopropyl)glycine). In group III, the mGluR4, mGluR6, mGluR7, and mGluR8 are linked to inhibition of cAMP formation by AC and respond effectively to L‐AP4 (L‐2‐amino‐4‐phosphonobutyrate; Schoepp et al., 1999). All mGluRs have a large hydrophilic N‐terminal domain (500–600 amino acids long) that forms the ligand‐binding region (> Figure 17‐2). The glutamate‐binding site is between two globular domains with a hinge region (Kunishima et al., 2000). X‐ray crystal structures of the ligand‐binding domain of mGluR1 with and without bound ligand have been reported, allowing conformational changes in the presence of glutamate and antagonists to be elucidated (Tsuchiya et al., 2002). The seven transmembrane domains are connected by short intra‐ and extracellular loops and an intracellular C‐terminal domain of variable length. The G‐protein‐coupling region corresponds to the intracellular loops separated by the transmembrane domains. The C terminus is involved in the targeting and tethering of mGluRs to specific neuronal compartments and interaction with mGluRs have been shown to form dimers, stabilized by disulfide bonds, which are thought to be involved in receptor activation (Romano et al., 1996a). Activation of mGluRs can modulate a variety of ion channels leading to changes in neuronal excitability. This is demonstrated in hippocampal neurons in which mGluR activation results in a reduction in leak Kþ conductance, the Ca2þ‐dependent slow after hyperpolarizing current and a slowly inactivating voltage‐dependent current termed IK(slow). These actions of mGluR activation lead to a dramatic increase in neuronal excitability (Conn and Pinn, 1997). Activation of mGluRs has also been shown to either increase or decrease the current through voltage‐sensitive Ca2þ channels depending on cell type. Activation of presynaptic mGluRs that serve as autoreceptors leads to a reduction of transmission at glutamatergic synapses. The mechanism of mGluR‐mediated reduction of synaptic transmission is not known. However, a reduction in voltage‐dependent Ca2þ currents may have a role. It should be noted that under certain conditions activation of mGluRs can lead to an increase in glutamate release. Presynaptic mGluRs have also been shown to reduce g‐aminobutyric acid (GABA) release, thereby reducing inhibitory transmission. Activation of mGluRs has also been shown to enhance NMDAR‐mediated responses (Conn and Pinn, 1997). In addition to ionotropic signaling, KARs can also enhance cortical excitability via second‐messenger cascades, an indirect regulation of ion channels through activation of G‐proteins (Lerma, 2003). While the molecular mechanism of this G‐protein interaction is not clear, postsynaptic KA‐2 subunits appear to be necessary for the metabotropic actions of KARs at hippocampal mossy fiber synapses (Ruiz et al., 2005).

3

Developmental Changes in Glutamate Receptors

3.1 Developmental Changes in AMPA Receptors The GluR1–3 subunits are detected in the CNS at embryonic day E15, and levels increase progressively during late embryonic and early postnatal days in the hippocampus and cerebral cortex but decreases in the striatum (Martin et al., 1998; To¨nnes et al., 1999). The GluR4 subunit is mainly expressed around late postnatal development and in adult brains. The properties of AMPARs undergo developmental changes as a result of the differential expression of alternatively spliced and edited subunits. In situ hybridization shows that flip forms of AMPAR subunits are expressed throughout embryonic and postnatal life and remain largely invariant during postnatal development. In contrast, flop forms arise at postnatal stages (Konig et al., 1992; To¨nnes et al., 1999). Receptors with flip sequences allow more current into cells than receptors containing flop sequences and this may be relevant to early‐forming synapses. GluR2 subunit Q/R editing has been shown to have consequences for brain development. During embryonic development, a small number of receptors appear to have the unedited GluR2 subunit, but with postnatal development virtually all (99%) GluR2 subunits are edited (Burnashev and Rozov, 2000). There is a developmental increase in the relative ratio of GluR2

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compared with other AMPAR subunits on the neuronal surface (Pickard et al., 2000). The relative increase in synaptic GluR2 reduces Ca2þ influx by forming Ca2þ‐impermeable AMPARs. These results indicate that not only the expression and synaptic targeting of AMPARs is regulated, but also the subunit composition is under developmental control. These findings are consistent with electrophysiological studies in brain slices, reporting changes in Ca2þ permeability consistent with the incorporation of GluR2 subunit‐containing AMPARs during development in neocortex (Kumar et al., 2002) and the activity‐dependent insertion of Ca2þ‐impermeable AMPARs at cerebellar stellate cell synapses (Liu and Cul‐Candy, 2000).

3.2 Developmental Changes in NMDA Receptors Glutamatergic synapses acquire NMDARs early in development. Subsequently, during the activity‐ dependent critical period, the subunit composition of NMDARs changes, altering their synaptic attachment and kinetic properties. While the NR1 subunit is ubiquitously present at a high level in virtually all rat brain regions at both embryonic and postnatal ages, the NR2A–D subunits are differentially distributed throughout the brain, with patterns of expression that change strikingly during development (Monyer et al., 1994; Watanabe, 1997). At birth, forebrain NMDARs are composed almost exclusively of NR1 and NR2B subunits, gradually incorporating more NR2A subunits during postnatal development. In the adult hippocampus and neocortex, NMDARs are mainly composed of NR1/NR2A/NR2B heteromers (Chazot and Stephenson, 1997; Hawkins et al., 1999), but the presence of other subunit combinations (e.g. homo‐ oligomeric complexes of NR1) have also been proposed (Garcia‐Gallo et al., 2001). Functional studies have examined the possible subunit composition of NMDARs in a number of identified neurons in various regions of the CNS (Cull‐Candy et al., 2001). There appears to be a general trend toward a decreasing (but still significant) contribution from the NR2B subunit during development, which is associated with an increased contribution of NR2A‐containing NMDARs to the synaptic current. These electrophysiological studies have reported that the duration of NMDAR‐mediated synaptic responses are shorter in older animals compared with younger ones. These changes correlate with the developmental switch in expression of NR2B subunit with slower deactivation kinetics to the NR2A subunit, which imparts faster deactivation kinetics (Cull‐Candy et al., 2001). Synaptic activity is a key factor in the regulation of NR2A and NR1b NMDAR subunit expression during development. A rapid and long‐lasting change in subunit composition has been identified for NMDARs in response to synaptic activity (Quinlan et al., 1999; Garcia‐Gallo et al., 2001) and experience (Quinlan et al., 1999), which suggests differential expression and synaptic targeting of various subunit combinations. The NR2C subunit appears postnatally and is only prominent in the cerebellum (Monyer et al., 1994; Watanabe, 1997). The NR2D subunit is mainly present in the diencephalon and brain stem at embryonic and neonatal stages (Monyer et al., 1994). The NR3 subunit is abundant within the late prenatal and early postnatal brain development (Sun et al., 1998). The changes in the kinetic properties and Ca2þ permeability of NMDARs are thought to be critical in the role that these receptors play in many activity‐dependent developmental processes. For example, the previously described developmental change to more NR2A subunits was reversed using transgenic mice, overexpressing the NR2B subunit (Tang et al., 1999). In the adult NR2B transgenic mice, glutamate evoked larger and longer NMDAR‐mediated currents and long‐term potentiation (LTP) was greatly enhanced (which was reminiscent of LTP in young rather than adult animals) probably due to the enhanced Ca2þ flux. The NR2B transgenic mice also showed improved learning scores compared with normal mice in different tests of their ability to acquire and retain information (Tang et al., 1999). These experiments suggest that the NMDARs serve as a graded molecular switch for gating the age‐dependent threshold for synaptic plasticity and memory formation.

3.3 Developmental Changes in Kainate Receptors The expression of KAR subunit transcripts and their mRNA editing change markedly during development. The mRNA for GluR5–7 and KA‐2 are detectable in E12 embryonic brain (Bahn et al., 1994).

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KAR subunits undergo a peak in their expression in the late embryonic and early postnatal period. GluR5 gene expression peaks around birth in the sensory cortex, in CA1 hippocampal interneurons in the stratum oriens, in the septum, and in the thalamus. GluR6 shows a prenatal expression peak in the cingulated gyrus of the neocortex. KA‐1 transcripts appear with the development of the hippocampus and remain largely confined to discrete areas such as the CA3 region, the dentate gyrus, and subiculum. KA‐2 transcripts are found throughout the CNS from as early as E12 and remain constant until adulthood (Bahn et al., 1994). Single‐cell RT‐PCR has shown that only GluR6 subunit mRNA is present in most of the cultured embryonic hippocampal neurons, exhibiting the characteristic native KAR response (Ruano et al., 1995). Cerebellar granule cells contain functional KARs both in situ and in culture, and mRNA editing of GluR5 and GluR6 increases, and KA‐2 expression decreases as granule cells mature (Smith et al., 1999). In contrast to GluR2, editing of the Q/R site in the M2 segment in GluR5 and GluR6 mRNAs (but not in GluR7) occurs at very low levels in embryonic brain and increases to 50% (GluR5) and 80% (GluR6) of the mRNA transcripts within the first few days after birth in most regions of the brain (Bernard et al., 1999). This developmentally regulated editing alters a critical residue producing subunits, which combine to form receptors with much lower Ca2þ permeabilities and single‐channel conductances. Changes in KAR Ca2þ permeability could be involved in the regulation of synapse formation, stabilization and plasticity (Dingledine et al., 1999).

3.4 Developmental Changes in Metabotropic Glutamate Receptors The expression of the mGluR subtypes is differentially regulated during development, displaying distinct regional and temporal profiles. The group I receptor subtypes, mGluR1 and mGluR5, are detected during embryonic development (Shigemoto et al., 1992; Lopez‐Bendito et al., 2002). mGluR1 expression gradually increases during the early postnatal days (Shigemoto et al., 1992; Lopez‐Bendito et al., 2002). mGluR1a predominates during development, whereas mGluR1b and mGluR1c appear in adulthood (Casabona et al., 1997). mGluR5 expression increases perinatally, reaching a peak in the second postnatal week and decreasing thereafter to adult levels (Catania et al., 1994; Romano et al., 1996b, 2002). The mGluR5a is the predominant splice variant during early development, with mGluR5b increasing subsequently and dominating in adulthood (Minakami et al., 1995; Romano et al., 1996b, 2002). A recent study of the perirhinal cortex indicates that the developmental reduction of mGluR5 was dependent on visual experience (Jo et al., 2006). Within group II, mGluR2 mRNA expression is low at birth and increases during postnatal development. In contrast, mGluR3 is highly expressed at birth and decreases during maturation (Catania et al., 1994). The group III mGluRs are also differentially expressed during development. mGluR4 expression is low at birth and increases during postnatal development (Catania et al., 1994; Elezgarai et al., 1999). mGluR7a levels are highest at P7 and P14, followed by a decline thereafter in cortical regions (Bradley et al., 1998).

4

Distribution of Glutamate Receptors in the CNS

Glutamate receptors are present in nearly all neurons and glial cells in the CNS. Each iGluR subunit and mGluR isoform shows a distinct pattern of distribution in different brain regions (Hollmann and Heinemann, 1994). For example, GluR2 and NR1 are both abundant and widespread (> Figure 17‐3). Others have a more restricted distribution and are abundant in some areas of the brain and populations of neurons. For example, GluR1 subunits are abundant in most neurons of the hippocampus and NR2C proteins are expressed in cerebellar granule cells (> Figure 17‐3).

4.1 Synaptic Distribution of Glutamate Receptors An important aspect of glutamatergic synaptic transmission is the recruitment and retention of iGluRs and mGluRs at their functional sites. The efficacy of synaptic transmission depends on appropriate targeting

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. Figure 17‐3 Regional distribution of iGluR subunits. AMPA, kainate, and NMDA receptor protein distribution was visualized on histoblots (To¨nnes et al., 1999) of adult rat brain horizontal sections using affinity‐purified anti‐GluR1–4 (Pickard et al., 2000), anti‐GluR1 (Molna´r et al., 1993), anti‐GluR1flop (To¨nnes et al., 1999), anti‐GluR2 (Chemicon International, Inc., Temecula, CA, USA), anti‐GluR4 (Baude et al., 1995), anti‐GluR5–7 (Gallyas et al., 2003), anti‐ GluR6/7 (Petralia et al., 1994a), anti‐KA‐2 (Molna´r et al., 1995), anti‐NR1 (Molna´r et al., 1995), anti‐NR2A, anti‐ NR2B, and anti‐NR2C (Chemicon International, Inc., Temecula, CA, USA) antibodies

and accumulation of glutamate receptors of appropriate number and type to the neuronal membrane near to the glutamate release sites. The glutamatergic synapses typically exhibit an electron‐dense postsynaptic density (PSD), where iGluRs are concentrated as shown by immunogold labeling (Baude et al., 1995; Nusser et al., 1998; Tanaka et al., 2005). Immunocytochemical and electrophysiological studies indicate that

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NMDARs are recruited first to glutamatergic synapses. Early in postnatal development, synapses acquire AMPARs with little change in NMDAR numbers (Golshani et al., 1998; Petralia et al., 1999; Pickard et al., 2000). With regard to their synaptic localization, AMPARs and NMDARs are concentrated at the PSD during postnatal development and adulthood (> Figure 17‐4; Baude et al., 1995; Nusser et al., 1998; Petralia et al., 1999; Takumi et al., 1999; Racca et al., 2000). In cortical neurons, NR2A NMDAR subunit has a

. Figure 17‐4 Schematic diagram illustrating the synaptic location of iGluRs and mGluRs. The combinations and precise localization of different iGluRs and mGluRs in synapses are brain region and cell‐type specific. Therefore, the schematic diagram is not representative of all glutamatergic synapse

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preferentially synaptic location (Stocca and Vicini, 1998). While the precise distribution of KARs at synapses remains to be determined, it is established that these receptors are located at synaptic sites in the adult CNS (Petralia et al., 1994a; Darstein et al., 2003). Immunohistochemical analysis of hippocampal mossy fiber synapses indicate a predominantly presynaptic localization for KA‐1 and greater postsynaptic concentration of KA‐2 subunit‐containing KARs (Darstein et al., 2003). Unlike iGluRs, mGluRs are not concentrated at the PSD (> Figure 17‐4). Immunohistochemical studies have demonstrated that group I mGluR1 and mGluR5 are excluded from the PSD and their highest concentration occurs perisynaptically (at the edge of the postsynaptic specialization), as well as along the extrasynaptic plasma membrane (> Figure 17‐4; Lujan et al., 1997; Lopez‐Bendito et al., 2002). Group II mGluR2 and mGluR3 can be found both at postsynaptic and presynaptic sites, depending on the brain region and cell type. At their postsynaptic location, mGluR2 is preferentially located outside the synapse (Lujan et al., 1997). In contrast, mGluR3 is found to be associated with glutamatergic synapses, including the postsynaptic specialization (Tamaru et al., 2001). At presynaptic locations, mGluR2 and mGluR3 are concentrated in the preterminal portion of axons or along the extrasynaptic membrane of axon terminals, and always outside the presynaptic active zone (Lujan et al., 1997; Shigemoto et al., 1997; Tamaru et al., 2001). Regarding group III mGluR4, mGluR7 and mGluR8 are mainly concentrated along the presynaptic active zone of glutamatergic synapses (Shigemoto et al., 1996, 1997) where they likely to act as autoreceptors.

4.2 Extrasynaptic Glutamate Receptors In recent year, it has become increasingly clear that extrasynaptic glutamate receptors participate in nonsynaptic communication between neurons (Vizi, 2000; Vizi and Mike, 2006). Under physiological conditions, at normal firing rate only a small proportion of released glutamate is able to leave the synaptic cleft and diffuse away due to effective neural and glial uptake systems (Danbolt, 2001). These glutamate transporters regulate the availability of released glutamate to pre‐ and postsynaptic receptors, shaping the response of the target cell and regulating the spillover of glutamate to neighboring synaptic sites, thereby maintaining the fidelity of synaptic transmission (Huang and Bergles, 2004). However, under certain conditions extrasynaptic glutamate spillover and nonsynaptic glutamatergic transmission still exists (Kullmann, 2000; Vizi, 2000; Vizi and Mike, 2006). A number of sources such as reversed uptake, glial exocytosis, osmotically driven release and neurotransmitter spillover can build up the extracellular concentration of glutamate, which can activate extrasynaptic glutamate receptors. In addition to synapses, AMPARs (Baude et al., 1995; Nusser et al., 1998), KARs (Petralia et al., 1994a; Kieval et al., 2001; Darstein et al., 2003), NMDARs (Petralia et al., 1994b, c), mGluR1 (Baude et al., 1993; Lujan et al., 1997), mGluR5 (Lujan et al., 1997; Hubert et al., 2001), and mGluR2/3 (Lujan et al., 1997; Shigemoto et al., 1997; Azkue et al., 2000) have been identified in the extrasynaptic plasma membrane (> Figure 17‐4). The lateral mobility and surface trafficking of AMPARs between synaptic and extrasynaptic sites in the plasma membrane are regulated by neuronal activity (Groc et al., 2004; Oh et al., 2006; Triller and Choquet, 2005). Extrasynaptic distribution of KARs, NMDARs, and mGluRs suggests some subunit composition and subtype specificity (Lujan et al., 1997; Stocca and Vicini, 1998; Massey et al., 2004; Jaskolski et al., 2005). For example, NR2B and NR2D subunit‐containing NMDARs (Stocca and Vicini, 1998; Lozovaya et al., 2004) and mGluR2 (Lujan et al., 1997) appear to be predominantly extrasynaptic. The affinity of predominantly extrasynaptic receptor isoforms toward glutamate is considerably higher compared with the synaptic counterparts (Vizi, 2000; Vizi and Mike, 2006). Furthermore, extrasynaptic receptors can be selectively targeted by drugs, due to their better accessibility and different pharmacological profile (Vizi, 2000; Vizi and Mike, 2006). In addition to neurons, astrocytes and oligodendrocytes also express functional iGluRs, mGluR3, and mGluR5 where these receptors are involved in neuronal to glial cell signaling (Winder and Conn, 1996; Luyt et al., 2003, 2006; Jensen, 2005). During development, iGluRs and mGluRs are transiently expressed in immature oligodendroglial cells which increase their vulnerability (Jensen, 2005; Luyt et al., 2006).

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Regulation of Glutamate Receptors during Synaptic Plasticity

The ability to regulate synaptic strength is critical feature of the CNS and is thought to underlie processes such as learning and memory. LTP and long‐term depression (LTD) have been studied intensively to understand the cellular and molecular mechanisms of synaptic plasticity (Bliss and Collingridge, 1993; Bear and Abraham, 1996). LTP is characterized by a long‐lasting increase in synaptic strength that is caused by a brief period of coordinated neuronal activity. Although LTP is persistent, the increase in synaptic strength can be reversed by different patterns of neuronal activity by a process known as depotentiation. These same patterns of neuronal activity can, under certain circumstances, lead to LTD of synaptic transmission from the basal (naı¨ve or nonpotentiated) state, and this de novo LTD can be reversed by a process of dedepression. Bidirectional and reversible alterations in synaptic efficiency make possible the dynamic storage of vast amount of neurally encoded information. Studies of LTP and LTD indicate that the activity and synaptic distribution of iGluRs and mGluRs are dynamically regulated and are fundamentally important for the short‐ and long‐term modification of synaptic efficacy. The molecular mechanisms that underlie these modulations of iGluRs and mGluRs include the interplay between different receptors, phosphorylation of specific sites and a wide range of receptor‐interacting proteins.

5.1 Phosphorylation of Glutamate Receptors and Synaptic Plasticity Phosphorylation represents a rapid and reversible way in which the function of proteins can be modified. Dynamic protein phosphorylation directly modulates iGluR and mGluR properties. Rapid and short‐term changes in phosphorylation can alter electrophysiological characteristics, protein–protein interactions and synaptic delivery or internalization of receptors. These changes underlie the major molecular mechanisms that affect many forms of synaptic plasticity.

5.1.1 Phosphorylation of AMPA Receptors AMPARs can be phosphorylated on their subunits GluR1, GluR2, and GluR4. All phosphorylation sites reside at serine (S), threonine (T), or tyrosine (Y) on the intracellular C‐terminal domain (> Figure 17‐2). Several key protein kinases, such as protein kinase A (PKA), protein kinase C (PKC), Ca2þ/calmodulin‐ dependent protein kinase II (CaMKII) and tyrosine kinases (Trks; receptor or nonreceptor family Trks), are involved in the site‐specific regulation of AMPAR phosphorylation. Other glutamate receptors (NMDARs, mGluRs) also regulate AMPARs through protein phosphorylation mechanisms. For example, the kinetics and magnitude of the NMDAR‐mediated Ca2þ signal determine the balance of phosporylation/dephosphorylation processes; a brief, large increase in Ca2þ favors the activation of kinases and longer, lower increase favors the activation of phosphatases (Lisman, 1989). In the C terminus, intracellular domains of GluR1‐S831 and GluR1‐S845 have been identified as major phosphorylation sites (> Figure 17‐5a). GluR1‐S831 is phosphorylated by PKC and CaMKII, whereas GluR1‐ S845 is phosphorylated by PKA (Roche et al., 1996; Mammen et al., 1997). Phosphorylation of these sites has been linked to the potentiation of AMPAR currents (Roche et al., 1996). Phosphorylation of the GluR1‐S831 site enhances single‐channel conductance (Derkach et al., 1999), whereas phosphorylation of the GluR1‐S845 residue confers increased open probability (Banke et al., 2000) and increased peak amplitude of the current (Roche et al., 1996) to the receptor channel. Site‐directed mutagenesis of the GluR2 C‐terminal domain identified GluR2‐S863 and GluR2‐S880 as PKC phosphorylation sites (Matsuda et al., 1999; McDonald et al., 2001). Phosphorylation of GluR2‐S880 modulates GluR2 interaction with glutamate receptor interacting protein‐1 (GRIP1) and protein interacting with C‐kinase‐1 (PICK1) (Chung et al., 2000). PKC phosphorylation of GluR2‐S880 is required for cerebellar LTD (Steinberg et al., 2006). In GluR4, a phosphorylation site for PKA, PKC, and CaMKII was found on S842 (Carvalho et al., 1999). GluR4‐T830 also appears to be phosphorylated by PKC (Carvalho et al., 1999). The amino acid residues situated in the vicinity of the S842 site on GluR4 are involved in an interaction with cytoplasmic proteins that prevent

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. Figure 17‐5 Schematic illustration of AMPAR phosphorylation sites and changes in synaptic plasticity. (a) Alignment of AMPAR C‐terminal domain amino acid sequences with known phosphorylation sites. Bold and underlined amino acids represent phosphorylation sites (marked by arrow heads). Phosphorylation of GluR1 subunit occurs at serine (S) residues S831 and S845, GluR2 subunits at S863, S880 and tyrosine (Y) Y876, and GluR4 at threonine (T) T830 and S845. (b) Some of the changes in AMPAR phosphorylation during hippocampal NMDAR‐ dependent LTP (NMDAR‐LTP), LTD (NMDAR‐LTD) and DHPG‐induced mGluR5‐dependent LTD (mGluR5‐LTD). In un‐potentiated naı¨ve synapses (middle panel) AMPARs are phosphorylated at GluR1‐S845 and GluR2 tyrosine residues. NMDA‐LTP (evoked by high‐frequency stimulation; HFS; top panel) is associated with CaMKII‐ mediated increased phosphorylation of GluR1‐S831. NMDA‐LTD (evoked by low‐frequency stimulation; LFS; bottom left panel) is coupled to increased phosphatase activity (including PP1/2A, 2B) and dephosphorylation of GluR1‐S845. Additionally, GluR2‐S880 phosphorylation by PKC disrupts the GluR2–GRIP interaction and triggers endocytosis following LTD induction (Seidenman et al., 2003). LFS given to a previously potentiated synapse (NMDAR‐LTP) dephosphorylates GluR1‐S831. HFS delivered to previously depressed synapses (NMDA‐ LTD) phosphorylates GluR1‐S845 (Lee et al., 2000). Activation of group I and group II mGluRs does not alter the phosphorylation state of GluR1‐S845 (Harris et al., 2004). Protein tyrosine phosphatases (PTP) are selectively involved in mGluR5‐LTD (bottom right panel; Moult et al., 2006)

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synaptic incorporation and that phosphorylation of the GluR4‐S842 site relieves this retention and is thus required for synaptic expression (Esteban et al., 2003). Despite the considerable sequence similarity with GluR2 (> Figure 17‐5a), the phosphorylation of GluR3 has not been reported yet. In addition to the phosphorylation on the serine and threonine residues, tyrosine phosphorylation of AMPARs occurs. The GluR2 C terminus has multiple tyrosine residues (> Figure 17‐5a). Src family Trks‐induced phosphorylation of GluR2‐Y876 has been reported (Hayashi and Huganir, 2004). Tyrosine phosphorylation at this site appears to inhibit the interaction of the GluR2 subunit with the PDZ domain‐containing proteins and GRIP1/2 but not PICK1 (Hayashi and Huganir, 2004).

5.1.2 Phosphorylation of Kainate Receptors GluR52b can be phosphorylated by PKC at residues S880 and S886 (Hirbec et al., 2003). Active PKC enhances the KAR currents and increases the retention of KARs at the synapse by interacting with anchoring proteins GRIP1/2 and PICK1 (Hirbec et al., 2003). The potentiation of KAR responses can be regulated by mGluR5 that links to PKC signaling (Cho et al., 2003). This process may include the serine phosphorylation of GluR52b. Recombinant GluR5 can also be phosphorylated by PKA in vitro (Cho et al., 2003), however the significance of this remains to be determined. PKA phosphorylation of recombinant GluR6 homomers can increase the open probability of the channel without significant changes in the time course of the response (Raymond et al., 1993; Wang et al., 1993). The previously identified residues (GluR6‐S684 and GluR6‐S666) are part of the extracellular M3–M4 linker region; therefore they are unlikely sites for phosphorylation by intracellular PKA (Raymond et al., 1993; Wang et al., 1993).

5.1.3 Phosphorylation of NMDA Receptors In the alternatively spliced intracellular C‐terminal C1‐cassette of NR1, PKC, and PKA phosphorylate S890 (PKC), S896 (PKC), and S897 (PKA) (Tingley et al., 1997). Phosphorylation of these sites regulates cell surface expression and clustering of NMDARs (Ehlers et al., 1995; Tingley et al., 1997; Crump et al., 2001; Fong et al., 2002; Scott et al., 2003) and may affect channel function by modulating the inhibitory interaction between NR1 and calmodulin (Ehlers et al., 1996; Hisatsune et al., 1997). NR1‐S890 and NR1‐ S896 are preferentially phosphorylated by PKCg and PKCa, respectively (Sa´nchez‐Pe´rez and Felipo, 2005). Activation of group I mGluRs by DHPG increases NR1‐S890 but not NR1‐S896 phosphorylation. Surface‐ expressed NR1 proteins are phosphorylated at S890 but not at S896 (Sa´nchez‐Pe´rez and Felipo, 2005). The C‐terminal domains of NR2A and NR2B are substrates for CaMKII (Omkumar et al., 1996), PKC, PKA (Leonard and Hell, 1997; Liao et al., 2001), and tyrosine kinases (Moon et al., 1994; Lau and Huganir, 1995). Several potential sites of tyrosine phosphorylation have been identified in NR2A and NR2B with Y1387 in NR2A and Y1472 in NR2B, representing major sites of phosphorylation by Src‐family kinases (Nakazawa et al., 2001; Yang and Leonard, 2001). Conversely, the NR1 intracellular C terminus does not appear to be tyrosine‐phosphorylated (Lau and Huganir, 1995). Tyrosine phosphorylation potentiates the NMDAR ion channel resulting in increased Ca2þ currents (Ali and Salter, 2001) and has been implicated in regulation of the internalization of NMDARs (Vissel et al., 2001; Li et al., 2002). In NR2C, S1244 is phosphorylated by both PKA and PKC (Chen et al., 2006). Phosphomimetic mutation of NR2C‐S1244 accelerates channel kinetics by increasing the speed of both the rise and decay of NMDA‐evoked currents (Chen et al., 2006). Increased PKA activity can facilitate the induction of LTP by increasing the Ca2þ permeability of NMDARs in dendritic spines (Skeberdis et al., 2006).

5.1.4 Phosphorylation of mGluRs The activity of several of the mGluRs has been shown to be modified via phosphorylation of the intracellular domains. The mGluR1c isoform has been shown to be phosphorylated by PKC (Ciruela et al., 1999). The phosphorylation of the mGluR1a isoform by both PKA and PKC is reported to regulate the

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intracellular signaling pathway of the receptor (Francesconi and Duvoisin, 2000; De Blasi et al., 2001). The phosphorylation of the receptor by PKC at mGluR1a‐T695, in the second intracellular loop, inhibits the agonist‐induced stimulation of the inositol 1,4,5‐trisphosphate (IP3) pathway. The phosphorylation of mGluR1a‐T695 is postulated to downregulate Gq/11 coupling to the receptor (Francesconi and Duvoisin, 2000). The phosphorylation of mGluR1a by PKA exerts the opposite effect in that it promotes via the IP3 signal transduction pathway (Francesconi and Duvoisin, 2000). The phosphorylation of mGluR5a and mGluR5b by PKC is responsible for the desensitization of the receptor (Gereau and Heinemann, 1998). Several sites were identified in point mutagenesis studies that were involved in the rapid desensitization of mGluR5: T606, S613 in the first intracellular loop, T665 in the second intracellular loop, and S881 and S890 in the C‐terminal domain (Gereau and Heinemann, 1998). The point mutation of one of several PKC phosphorylation sites resulted in a loss or a reduction in PLC coupling of the receptor (Gereau and Heinemann, 1998). PKC‐dependent phosphorylation of mGluR5‐S839 is required for the characteristic intracellular Ca2þ oscillations produced by mGluR5 activation (Kim et al., 2005). mGluR5 may also be phosphorylated by PKA, which could regulate the activity of the receptor (Cho et al., 2002). The activity of group III receptors is also modulated by the phosphorylation by PKC (Nakajima et al., 1999) at mGluR7a‐S862 on the C‐terminal domain (Airas et al., 2001). The phosphorylation of mGluR7a has been shown to be negatively regulated by PICK1 in vitro (Dev et al., 2000). The mGluR7a subtype is located presynaptically where it regulates feedback inhibition of glutamate release from the presynaptic terminus via a Ca2þ/calmodulin‐dependent mechanism. The binding of calmodulin to the C‐terminal domain of mGluR7a has been postulated to enable the release of G‐protein binding, enabling the mediation of this autoinhibition (O’Connor et al., 1999). The phosphorylation of mGluR7a‐S862 disrupts the binding of calmodulin and thereby modulates the functional properties of the receptor (Nakajima et al., 1999; Airas et al., 2001).

5.2 Receptor Trafficking and Synaptic Plasticity Glutamate receptors have been shown to move rapidly around the neuron to alter the number and molecular composition of receptors that are available to respond to released neurotransmitters (Collingridge et al., 2004). The receptors are inserted into and removed from the plasma membrane by exocytosis and endocytosis, respectively, and hence diffuse laterally within the plasma membrane. While the molecular mechanisms regulating the differential targeting and clustering of different iGluRs and mGluRs in vivo are unclear, proteins that interact directly or indirectly with glutamate receptors are likely to play central roles in this process. This section describes examples of rapid trafficking of glutamate receptors and how this relates to our understanding of the molecular basis of synaptic plasticity.

5.2.1 Recruitment of AMPA Receptors at Synapses Studies of AMPAR targeting in synaptic plasticity have focused on the idea that alteration in AMPAR number is one of the expression mechanisms for LTP and LTD (> Figure 17‐6). Now it is generally accepted that AMPARs are inserted into the membrane during NMDAR‐dependent LTP (Lu et al., 2001; Pickard et al., 2001) and they are internalized during NMDA‐LTD (Beattie et al., 2000; Carroll et al., 2001) and mGluR‐LTD (Snyder et al., 2001; Xiao et al., 2001). AMPARs can also move laterally within the plasma membrane (Triller and Choquet, 2005). After exocytosis, AMPARs are diffusely distributed along dendrites before accumulating at synaptic sites (Passafaro et al., 2001). In NMDA‐LTD, AMPARs first diffuse from synaptic to extrasynaptic sites where they are internalized (Ashby et al., 2004). A general model is emerging in which synaptic plasticity involves both regulated exocytosis and endocytosis of AMPARs at extrasynaptic sites and their regulated lateral diffusion into and out of the synapse (Collingridge et al., 2004). Proteins that directly bind to AMPAR subunits are likely regulators of rapid AMPAR trafficking at synapses (> Figure 17‐6; Bredt and Nicoll, 2003; Collingridge et al., 2004). In NMDAR‐LTP, activation of CaMKII causes insertion of GluR1‐containing AMPARs involving the interaction with stargazing and

Molecular organization and regulation of glutamate receptors in developing and adult mammalian CNS . Figure 17‐6 Schematic diagram of different steps of AMPAR targeting. See > Section 5.2.1 for details

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synapse‐associated protein‐97 (SAP97), which binds to motor protein such as myosin VI (MyoVI; > Figure 17‐6; Wu et al., 2002). AMPARs are probably inserted at sites distant from the PSD, from which they diffuse laterally within the membrane (> Figure 17‐6; Triller and Choquet, 2005). A widely held hypothesis for principal neuron AMPAR trafficking in NMDAR‐LTP is that AMPARs composed of GluR1 and GluR2 subunits are incorporated at synapses during LTP and are subsequently replaced by AMPARs‐containing GluR2 and GluR3 subunits through constitutive recycling (Shi et al., 2001). However, a more recent study suggests that NMDAR‐LTP in CA1 hippocampal pyramidal neurons causes rapid incorporation of GluR2‐lacking Ca2þ‐permeable AMPARs (Plant et al., 2006). These Ca2þ‐permeable AMPARs are replaced by GluR2‐containing, Ca2þ‐impermeable AMPARs 25 min after LTP induction (Plant et al., 2006). One possible interpretation of these results is that Ca2þ‐permeable AMPARs may ‘‘tag’’ newly potentiated synapses by initiating Ca2þ‐dependent changes in protein synthesis, gene expression required for a stable increase in synaptic strength. Interesting to point out that during early development GluR2 lacking, Ca2þ‐permeable AMPARs appear first, which are replaced by GluR2‐containing, Ca2þ‐ impermeable AMPARs (see > Section 3.1; Pickard et al., 2000). This suggests that LTP and early postnatal development are mechanistically related and the sequential appearance of receptors with different ion selectivity and kinetic properties is likely to play an important role in changes of synaptic efficacy. NMDAR activation induces the translocation of CaMKII to the synapse, where it can directly interact with the NR2B NMDAR subunit, which locks CaMKII in an active conformation and enhances synaptic activity by phosphorylation and recruitment of AMPARs and other proteins (Barria and Malinow, 2005). Stargazin (and similar transmembrane AMPAR regulatory proteins) might help to concentrate AMPARs at the PSD by its interaction with the PDZ interaction site containing protein PSD‐95 (PSD protein‐95; > Figure 17‐6; Bredt and Nicoll, 2003; Nicoll et al., 2006). Interaction of AMPARs with phosphatidylinositol 3‐kinase (PI3K) also promote surface expression (Man et al., 2003). Interaction of GluR1 with protein 4.1 (4.1N) appears to stabilize AMPARs at the cell surface by cross‐linking them to actin cytoskeleton (> Figure 17‐6; Shen et al., 2000). PICK1 acts to downregulate the GluR2 content of AMPARs at hippocampal CA1 synapses, thereby increasing synaptic strength at resting membrane potentials (Terashima et al., 2004). PICK1 is also a Ca2þ sensor; stimulated Ca2þ influx enhances PICK1–GluR2 binding to initiate AMPAR endocytosis (Hanley and Henley, 2005). GluR2‐containing receptors are constitutively recycled involving the N‐ethylmaleimide‐sensitive factor (NSF). NSF is an ATPase that is involved in membrane fusion events and binds to the C‐terminal domain of GluR2 (Nishimune et al., 1998; Osten et al., 1998). The association between NSF and GluR2 is important to maintain AMPARs at synapses and may dissociate them from tethers, the multiple PDZ proteins GRIP and AMPAR‐binding protein (ABP; > Figure 17‐6). GRIP and ABP probably anchor AMPARs at both synaptic and intracellular locations (> Figure 17‐6; Daw et al., 2000; Osten et al., 2000). Disruption of the GluR2–NSF interaction leads to reduced surface expression of AMPARs (Luscher et al., 1999; Noe¨l et al., 1999). Following appropriate synaptic stimulation, the GluR2 interaction with NSF is exchanged to interaction with AP2 (an adapter complex binds to GluR2), which initiates clathrin‐dependent internalization (> Figure 17‐6; Lee et al., 2002). PICK1 can also bind the C‐terminal domain of GluR2 and target PKCa to phosphorylate GluR2‐S880. Once phosphorylated at S880, GluR2 can bind PIC1 but not GRIP/ABP, which provides a mechanism by which AMPARs can be freed from GRIP/ABP for internalization (Matsuda et al., 1999; Chung et al., 2000). Once untethered, AMPARs probably diffuse laterally in the plasma membrane to sites of endocytosis at the periphery of synapses (> Figure 17‐6; Ashby et al., 2004). Internalized AMPARs might be recycled to the membrane, held in an intracellular pool or degraded (> Figure 17‐6). The intracellular pool is also probably tethered by PDZ proteins such as GRIP/ABP, and can be released by PICK1/PKCa(Daw et al., 2000). The synaptic/extrasynaptic redistribution of AMPARs and their lateral movement in the neuronal plasma membrane are also regulated processes. For example, GluR1‐S845 phosphorylation can traffic AMPARs to extrasynaptic sites for subsequent delivery to synapses during NMDAR‐LTP (Sun et al., 2005; Oh et al., 2006).

5.2.2 Kainate Receptor Trafficking Cell surface KARs are clustered by PSD‐95 and related SAP family members (Garcia et al., 1998). The cell surface expression of KARs1 is determined by the subunit composition and alternative splicing of receptors. Low level of GluR5a and GluR5b are present on the cell surface, whereas GluR5c is retained in the

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endoplasmic reticulum (ER). Small amounts of GluR6b are present at the cell surface but GluR6a is delivered efficiently to the plasma membrane (Jaskolski et al., 2005). KA‐2 is restricted to the ER (Gallyas et al., 2003; Ren et al., 2003; Nasu‐Nishimura et al., 2006). When assembled in heteromeric receptors, GluR6a promotes the surface expression of all ER‐retained subunits. Disrupting interactions of GluR5 and GluR6 with GRIP or PICK1 leads to a rapid reduction in KAR‐mediated synaptic transmission (Hirbec et al., 2003). Considering the role of GRIP and PICK1 in AMPAR targeting, it is likely that these interaction partners also involved in KAR trafficking. GluR6‐containing KARs are subjected to activity‐dependent endocytic sorting (Martin and Henley, 2004). Kainate activation of hippocampal neuronal cultures causes a PKC‐dependent internalization of KARs that are targeted to lysosomes for degradation. In contrast, NMDAR activation evokes a Ca2þ‐, PKA‐, and PKC‐dependent endocytosis of KARs to early endosomes with subsequent reinsertion back into the plasma membrane (Martin and Henley, 2004).

5.2.3 NMDA Receptor Trafficking NMDA‐mediated synaptic transmission can itself be modulated during synaptic plasticity, in a process termed metaplasticity. The regulated trafficking of NMDARs can fundamentally alter glutamatergic neurotransmission and the induction of several forms of synaptic plasticity. Exit from the ER represents a critical regulatory step in NMDAR trafficking. Neuronal cell surface targeting of NMDARs from intracellular compartments depends on subunit composition and assembly (McIlhinney et al., 1996; Wenthold et al., 2003). NR2 subunits are not transported to the cell surface unless they associate with NR1 subunits (McIlhinney et al., 1998). NR1 subunits do not form functional receptors alone, but their cell surface expression depends on the splice variant (Wenthold et al., 2003). This is due to the presence of ER retention/retrival motif in the C1 alternatively expressed cassette of the C‐terminal domain of NR1 (Standley et al., 2000). ER retention can be relieved by interaction with PDZ proteins, which facilitate surface expression (Standley et al., 2000). Subunit‐specific processes determine the subcellular distribution of NMDARs; NR1/NR2B‐containing NMDARs are constitutively inserted at synapses early in development and later replaced by NR1/NR2A receptors in an activity‐dependent manner (Barria and Malinow, 2002). At synapses, NMDARs are stabilized by interaction with PSD‐95 (Roche et al., 2001; Li et al., 2003). Surface expression of NMDARs can also be regulated by insulin receptor‐mediated tyrosine phosphorylation (Skeberdis et al., 2001) and by PKC (Lan et al., 2001a). PKC‐mediated insertion of NMDARs might be triggered by activation of mGluRs and involves a process that depends on synaptosomal‐associated protein‐25 (SAP25) (Lan et al., 2001b). An increase in NMDAR surface expression has been reported after the induction of LTP at adult CA1 synapses (Grosshans et al., 2002). NMDARs can be rapidly internalized in response to the binding of glutamate or glycine (Roche et al., 2001; Li et al., 2003; Nong et al., 2003). The association of NMDARs with PSD‐95 and their subsequent endocytosis are regulated by tyrosine phosphorylation. Phosphorylation of NR2B‐Y1472 interferes with binding to PSD‐95 and promotes clathrin‐dependent endocytosis, probably by promoting the binding of AP2 to NR2B. Dephosphorylation of NR1‐Y837 and NR2A‐Y842 might affect AP2 binding, promoting clathrin‐dependent endocytosis in a similar way (Vissel et al., 2001). A form of LTD that is induced by mGluR activation is associated with the internalization of NMDARs (Snyder et al., 2001). The activity‐dependent synaptic removal and rapid endocytosis of the developmentally regulated NR3A subunit‐containing NMDARs are regulated by PACSIN1/syndapin1 (Pe´rez‐Otan˜o et al., 2006). The NR3A interaction partner PACSIN1/ syndapin1 is a neuron‐specific multivalent adaptor that coordinates actin remodeling with cargo recruitment and membrane fission during endocytosis and assembles a complex of proteins including dynamin and clathrin (Pe´rez‐Otan˜o et al., 2006). By recruiting the endocytic machinery, NR3A may behave as an endocytic adaptor for the NMDAR itself, and its regulated synaptic removal could direct synapse elimination. There are mobile and immobile pools of NMDARs at synaptic and extrasynaptic sites (Groc et al., 2004). NMDARs can move from extrasynaptic to synaptic sites on the cell surface, so the number of NMDARs at synapses might be rapidly altered by lateral diffusion in the plasma membrane (Tovar and Westbrook, 2002; Groc et al., 2004). This process could be regulated by the activation of PKC (Groc et al., 2004). NMDAR subunit proteins are also trafficked within dendrites by a process that involves microtubules (Washbourne et al., 2002; Guillaud et al., 2003). Intradendritic trafficking of NR2B subunits requires them

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to associate with the motor protein KIF17 through LIN10 (Guillaud et al., 2003). Within the PSD, NMDARs are linked by a‐actin and spectrin to F‐actin, which associates with myosin and might be dynamically regulated by myosin light‐chain kinase (MLCK; Lei et al., 2001). Synaptic NR2A‐containing NMDARs seem to be important for LTP and extrasynaptic NR2B‐containing receptors are involved in LTD (Cull‐Candy et al., 2001). In adult neocortical slices, de novo LTD induction is enhanced by blockage of glutamate uptake, indicating that the diffusion of glutamate to extrasynaptic NR2B‐containing NMDARs triggers LTD (Massey et al., 2004). Furthermore, LTD can be induced after blockade of synaptic NMDARs (Massey et al., 2004). The activity‐dependent internalization of short C‐terminal GluR2‐containing AMPARs is selectively mediated by NR2B‐containing NMDARs (Tigaret et al., 2006). Therefore, NMDAR localization and targeting are crucial determinants of synaptic plasticity.

5.2.4 Trafficking of mGluRs Like other G‐protein‐coupled receptors, the surface distribution of mGluRs can be rapidly altered (Dhami and Ferguson, 2006). Various proteins interact with mGluRs and might regulate their trafficking in response to activity (Fagni et al., 2004). Agonist application causes rapid internalization of group I mGluRs through an arrestin‐ and dynamin‐dependent process (Dhami and Ferguson, 2006). In addition, mGluR5s are highly mobile in the lateral plane of the neuronal membrane, and binding to the scaffolding protein Homer stabilizes clusters of these receptors, whereas activation by an agonist increases the mobility of both clustered and nonclustered mGluR5 (Serge et al., 2002). Splice variants of Homer‐1 differentially regulate the trafficking of group I mGluRs to the plasma membrane (Roche et al., 1999; Ango et al., 2001). The regulated cell surface trafficking of mGluRs is likely to influence synaptic plasticity.

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Conclusions

Recent insight into how iGluRs and mGluRs are modified during development and rapidly reorganized at synapses has profound implications for our understanding of the mechanisms of neuronal plasticity and possible mechanisms of information storage in the brain. Complex interplay between different glutamate receptors and synaptic scaffolding proteins regulate the synaptic distribution and molecular composition of iGluRs and mGluRs. This process is dynamically regulated in response to neuronal activity, and thereby regulates synaptic plasticity. The challenge is to understand the spatiotemporal features of various phosphorylation events and specific protein–protein interactions and their relative importance during different forms of experience‐driven synaptic plasticity.

Acknowledgment The author is grateful to the Medical Research Council and the Wellcome Trust for financial support.

References Airas JM, Betz H, Far E. 2001. PKC phosphorylation of a conserved serine residue in the C‐terminus of group III metabotropic glutamate receptors inhibits calmodulin binding. FEBS Lett 494: 60-63. Ali DW, Salter MW. 2001. NMDA receptor regulation by Src kinase signalling in excitatory synaptic transmission and plasticity. Curr Opin Neurobiol 11: 336-342.

Ango F, Prezeau L, Muller T, Tu JC, Xiao B, et al. 2001. Agonist‐ independent activation of metabotropic glutamate receptors by the intracellular protein Homer. Nature 411: 962-965. Ashby MC, De La Rue Sa, Ralph GS, Uney J, Collingridge GL, et al. 2004. Removal of AMPA receptors (AMPARs) from synapses is preceded by transient endocytosis of extrasynaptic AMPARs. J Neurosci 24: 5172-5176.

Molecular organization and regulation of glutamate receptors in developing and adult mammalian CNS Azkue JJ, Mateos JM, Elezgarai I, Benitez R, Osorio A, et al. 2000. The metabotropic glutamate receptor subtype mGluR2/3 is located at extrasynaptic loci in rat spinal dorsal horn synapses. Neurosci Lett 287: 236-238. Bahn S, Volk B, Wisden W. 1994. Kainate receptor gene expression in the developing rat brain. J Neurosci 14: 5525-5547. Banke TG, Bowie D, Lee H, Huganir RL, Schousboe A, et al. 2000. Control of GluR1 AMPA receptor function by cAMP‐ dependent protein kinase. J Neurosci 20: 89-102. Barria A, Malinow R. 2002. Subunit‐specific NMDA receptor trafficking to synapses. Neuron 35: 345-353. Barria A, Malinow R. 2005. NMDA receptor subunit composition controls synaptic plasticity by regulating binding to CaMKII. Neuron 48: 289-301. Baude A, Nusser Z, Molna´r E, McIlhinney RAJ, Somogyi P. 1995. High‐resolution immunogold localization of AMPA type glutamate receptor subunits at synaptic and non‐ synaptic sites in rat hippocampus. Neuroscience 69: 1031-1055. Baude A, Nusser Z, Roberts JD, Mulvihill E, McIlhinney RAJ, et al. 1993. The metabotropic glutamate receptor (mGluR1 alpha) is concentrated at perisynaptic membrane of neuronal subpopulations as detected by immunogold reaction. Neuron 11: 771-787. Bear MF, Abraham WC. 1996. Long‐term depression in hippocampus. Annu Rev Neurosci 19: 437-462. Beattie EC, Carroll RC, Yu X, Morishita W, Yasuda H, et al. 2000. Regulation of AMPA receptor endocytosis by a signalling mechanism shared with LTD. Nat Neurosci 3: 1291-1300. Bernard A, Ferhat L, Dessi F, Charton G, Represa A, et al. 1999. Q/R editing of the rat GluR5 and GluR6 kainate receptors in vivo and in vitro: Evidence for independent developmental, pathological and cellular regulation. Eur J Neurosci 11: 604-616. Bliss TV, Collingridge GL. 1993. Asynaptic model of memory: Long‐term potentiation in the hippocampus. Nature 361: 31-39. Bradley SR, Rees HD, Yi H, Levey AI, Conn PJ. 1998. Distribution and developmental regulation of metabotropic glutamate receptor 7a in rat brain. J Neurochem 71: 636-645. Bredt DS, Nicoll RA. 2003. AMPA receptor trafficking at excitatory synapses. Neuron 40: 361-379. Burnashev N, Rozov A. 2000. Genomic control of receptor function. Cell Mol Life Sci 57: 1499-507. Carroll RC, Beattie EC, Zastrow von M, Malenka RC. 2001. Role of AMPA receptor endocytosis in synaptic plasticity. Nat Rev Neurosci 2: 315-324. Carvalho AL, Kameyama K, Huganir RL. 1999. Characterization of phosphorylation sites on the glutamate receptor 4 subunit of the AMPA receptors. J. Neurosci 19: 4748-4754.

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Casabona G, Knopfel T, Kuhn R, Gasparini F, Baumann P, et al. 1997. Expression and coupling to polyphosphoinositide hydrolysis of group I metabotropic glutamate receptors in early postnatal and adult rat brain. Eur J Neurosci 9: 12-17. Catania MV, Landwehrmeyer GB, Testa CM, Standaert DG, Penney JB Jr., et al. 1994. Metabotropic glutamate receptors are differentially regulated during development. Neuroscience 61: 481-495. Chazot PL, Stephenson FA. 1997. Molecular dissection of native mammalian forebrain NMDA receptors containing the NR1 C2 exon: Direct demonstration of NMDA receptors comprising NR1, NR2A, and NR2B subunits within the same cortex. J Neurochem 69: 2138-2144. Chen BS, Braud S, Badger JD II, Isaac JTR, Roche KW. 2006. Regulation of NR1/NR2C N‐methyl‐D‐aspartate (NMDA) receptors by phosphorylation. J Biol Chem 281: 16583-16590. Cho K, Brown MW, Bashir ZI. 2002. Mechanisms and physiological role of enhancement of mGluR5 receptor function by group II mGlu receptor activation in rat perirhinal cortex. J Physiol 540: 895-906. Cho K, Francis JC, Hirbec H, Dev K, Brown MW, et al. 2003. Regulation of kainate receptors by protein kinase C and metabotropic glutamate receptors. J Physiol 548(3): 723-730. Chung HJ, Xia J, Scannevin RH, Zhang X, Huganir RL. 2000. Phosphorylation of the AMPA receptor subunit GluR2 differentially regulates its interaction with PDZ domain‐ containing proteins. J Neurosci 20: 7258-7267. Ciruela F, Giacometti A, McIlhinney RA. 1999. Functional regulation of metabotropic glutamate receptor type 1c: A role for phosphorylation in the desensitization of the receptor. FEBS Lett 462: 278-282. Collingridge GL, Isaac JT, Wang YT. 2004. Receptor trafficking and synaptic plasticity. Nat Rev Neurosci 5: 952-962. Conn PJ, Pinn JP. 1997. Pharmacology and functions of metabotropic glutamate receptors. Annu Rev Pharmacol Toxicol 37: 205-237. Crump FT, Dillman KS, Craig AM. 2001. cAMP‐dependent protein kinase mediates activity‐regulated synaptic targeting of NMDA receptors. J Neurosci 21: 5079-5088. Cull‐Candy S, Brickley S, Farrat M. 2001. NMDA receptor subunits: Diversity, development and disease. Curr Opin Neurobiol 11: 327-335. Curtis DR, Phillis JW, Watkins JC. 1959. Chemical excitation of spinal neurones. Nature 183: 611-612. Danbolt NC. 2001. Glutamate uptake. Prog Neurobiol 65: 1-105. Darstein M, Petralia RS, Swanson GT, Wenthold RJ, Heinemann SF. 2003. Distribution of kainate receptor subunits at hippocampal mossy fiber synapses. J Neurosci 23: 8013-8019.

435

436

17

Molecular organization and regulation of glutamate receptors in developing and adult mammalian CNS

Daw MI, Chittajallu R, Bortolotto ZA, Dev KK, Duprat F, et al. 2000. PDZ proteins interacting with C‐terminal GluR2/3 are involved in a PKC‐dependent regulation of AMPA receptors at hippocampal synapses. Neuron 28: 873-886. De Blasi A, Conn PJ, Pin J, Nicoletti F. 2001. Molecular determinants of metabotropic glutamate receptor signalling. Trends Pharmacol Sci 22: 114-120. Derkach V, Barria A, Soderling TR. 1999. Ca2þ/calmodulin‐ kinase II enhances channel conductance of alpha‐amino‐3‐ hydroxy‐5‐methyl‐4‐isoxazolepropionate type glutamate receptors. Proc Natl Acad Sci USA 96: 3269-3274. Dev KK, Nakajima Y, Kitano J, Braithwaite SP, Henley JM, et al. 2000. PICK1 interacts with and regulates PKC phosphorylation of mGLUR7. J Neurosci 20: 7252-7257. Dhami G, Ferguson SSG. 2006. Regulation of metabotropic glutamate receptor signalling desensitization and endocytosis. Pharmacol Ther 111: 260-271. Dingledine R, Borges K, Bowie D, Traynelis SF. 1999. The glutamate receptor ion channels. Pharmacol Rev 51: 7-61. Ehlers MD, Tingley WG, Huganir RL. 1995. Regulated subcellular distribution of the NR1 subunit of the NMDA receptor. Science 269: 1734-1737. Ehlers MD, Zhang S, Bernhadt JP, Huganir RL. 1996. Inactivation of NMDA receptors by direct interaction of calmodulin with the NR1 subunit. Cell 84: 745-755. Elezgarai I, Benitez R, Mateos JM, Lazaro E, Osorio A, et al. 1999. Developmental expression of the group III metabotropic glutamate receptor mGluR4a in the medial nucleus of the trapezoid body of the rat. J Comp Neurol 411: 431-440. Esteban JA, Shi SH, Wilson C, Nuriya M, Huganir RL, et al. 2003. PKA phosphorylation of AMPA receptor subunits controls synaptic trafficking underlying plasticity. Nat Neurosci 6: 136-143. Fagni L, Ango F, Perroy J, Bockaert J. 2004. Identification and functional roles of metabotropic glutamate receptor‐ interacting proteins. Sem Cell Dev Biol 15: 289-298. Fong DK, Rao A, Crump FT, Craig AM. 2002. Rapid synaptic remodeling by protein kinase C: Reciprocal translocation of NMDA receptors and calcium/calmodulin‐dependent kinase II. J Neurosci 22: 2153-2164. Francesconi A, Duvoisin RM. 2000. Opposing effects of protein kinase C and protein kinase A on metabotropic glutamate receptor signaling: Selective desensitization of the inositol triphosphate/Ca pathway by phosphorylation of the receptor–G protein‐coupling domain. Proc Natl Acad Sci USA 97: 6185-6190. Gallyas F Jr., Ball SM, Molna´r E. 2003. Assembly and cell surface expression of KA‐2 subunit‐containing kainate receptors. J Neurochem 86: 1414-1427. Garcia EP, Mehta S, Blair LA, Wells DG, Shang J, et al. 1998. SAP90 binds and clusters kainate receptors causing incomplete desensitization. Neuron 21: 727-739.

Garcia‐Gallo M, Renart J, Diaz‐Guerra M. 2001. The NR1 subunit of the N‐methyl‐D‐aspartate receptor can be effectively expressed alone in the cell surface of mammalian cells and is required for the transport of the NR2A subunit. Biochem J 356: 539-547. Gereau RW, Heinemann SF. 1998. Role of protein kinase C phosphorylation in rapid desensitization of metabotropic glutamate receptor 5. Neuron 20: 143-151. Golshani P, Warren RA, Jones EG. 1998. Progression of change in NMDA, non‐NMDA, and metabotropic glutamate receptor function at the developing corticothalamic synapse. J Neurophysiol 80: 143-154. Groc L, Heine M, Cognet L, Brickley K, Stephenson FA, et al. 2004. Differential activity‐dependent regulation of the lateral mobilities of AMPA and NMDA receptors. Nat Neurosci 7: 695-696. Grosshans DR, Clayton DA, Coultrap SJ, Browning MD. 2002. LTP leads to rapid surface expression of NMDA but not AMPA receptors in adult rat CA1. Nat Neurosci 5: 27-33. Guillaud L, Setou M, Hirokawa N. 2003. KIF17 dynamics and regulation of NR2B trafficking in hippocampal neurons. J Neurosci 23: 131-140. Hanley JG, Henley JM. 2005. PICK1 is a calcium‐sensor for NMDA‐induced AMPA receptor trafficking. EMBO J 24: 3266-3278. Harris SL, Gallyas F, Molna´r E. 2004. Activation of metabotropic glutamate receptors does not alter the phosphorylation state of GluR1 AMPA receptor subunit at serine 845 in perirhinal cortical neurons. Neurosci Lett 372: 132-136. Hawkins LM, Chazot PL, Stephenson FA. 1999. Biochemical evidence for the co‐association of three N‐methyl‐D‐ aspartate (NMDA) R2 subunits in recombinant NMDA receptors. J Biol Chem 274: 27211-27218. Hayashi T. 1954. Effects of sodium glutamate on the nervous system. Keio J Med 3: 192-193. Hayashi T, Huganir RL. 2004. Tyrosine phosphorylation and regulation of the AMPA receptor by SRC family tyrosine kinases. J Neurosci 24: 6152-6160. Hermans E, Challiss RAJ. 2001. Structural, signalling and regulatory properties of the group I metabotropic glutamate receptors: Prototypic family C G‐protein‐coupled receptors. Biochem J 359: 465-484. Hirbee H, Francis JC, Lauri SE, Braithwaite SP, Coussen F, et al. 2003. Rapid and differential regulation of AMPA and kainate receptors at hippocampal mossy fibre synapses by PICK1 and GRIP. Neuron 37: 625-638. Hisatsune C, Umemori H, Inoue T, Michikawa T, Kohda K, et al. 1997. Phosphorylation‐dependent regulation of N‐methyl‐D‐aspartate receptors by calmodulin. J Biol Chem 272: 20805-20810.

Molecular organization and regulation of glutamate receptors in developing and adult mammalian CNS Hollmann M. 1999. Molecular structure of ionotropic glutamate receptors. Handbook of Experimental Pharmacology. Ionotropic Glutamate Receptors in the CNS, Vol. 141. Jonas P Monyer H, editors., Heidelberg: Springer. Hollmann M, Heinemann S. 1994. Cloned glutamate receptors. Ann Rev Neurosci 17: 31-108. Hollmann M, Maron C, Heinemann S. 1994. N‐glycosylation site tagging suggests a three transmembrane domain topology for the glutamate receptor GluR1. Neuron 13: 1331-1343. Hollmann M, O’Shea‐Greenfield A, Rogers SW, Heinemann S. 1989. Cloning by functional expression of a member of the glutamate receptor family. Nature 342: 643-648. Houamed KM, Kuijper JL, Gilbert TL, Haldeman BA, O’Hara PJ, et al. 1991. Cloning, expression, and gene structure of a G‐protein‐coupled glutamate receptor from rat brain. Science 252: 1318-1321. Huang YH, Bergles DE. 2004. Glutamate transporters bring competition to the synapse. Curr Opin Neurobiol 14: 346-352. Hubert GW, Paquet M, Smith Y. 2001. Differential subcellular localization of mGluR1a and mGluR5 in the rat and monkey substantia nigra. J Neurosci 21: 1838-1847. Jaskolski F, Coussen F, Mulle C. 2005. Subcellular localization and trafficking of kainate receptors. Trends Pharmacol Sci 26: 20-26. Jensen FE. 2005. Role of glutamate receptors in periventricular leucomalacia. J Child Neurol 20: 950-959. Jo J, Ball SM, Seok H, Oh SB, Massey PV, et al. 2006. Experience‐dependent modification of mechanisms of long‐term depression. Nat Neurosci 9: 170-172. Kieval JZ, Hubert GW, Charara A, Pare JF, Smith Y. 2001. Subcellular and subsynaptic localization of presynaptic and postsynaptic kainate receptor subunits in the monkey striatum. J Neurosci 21: 8746-8757. Kim CH, Braud S, Isaac JTR, Roche KW. 2005. Protein kinase C phosphorylation of the metabotropic glutamate receptor mGluR5 on serine 839 regulates Ca2þ oscillations. J Biol Chem 280(27): 25409-25415. Konig N, Drian MJ, Manzoni O, Sladeczeck F. 1992. Early functional glutamate receptors in acutely dissociated embryonic raphe cells. Neuroreport 3: 738-740. Kullmann DM. 2000. Spillover and synaptic cross talk mediated by glutamate and GABA in the mammalian brain. Prog Brain Res 125: 339-351. Kumar SS, Bacci A, Kharazia V, Huguenard JR. 2002. A developmental switch of AMPA receptor subunits in neocortical pyramidal neurons. J Neurosci 22: 30053015. Kunishima N, Shimada Y, Tsuji Y, Sato T, Yamamoto M, et al. 2000. Structural basis of glutamate recognition by a dimeric metabotropic glutamate receptor. Nature 407: 971-977.

17

Lan JY, Skeberdis VA, Jover T, Grooms SY, Lin Y, et al. 2001a. Protein kinase C modulates NMDA receptor trafficking and gating. Nat Neurosci 4: 382-390. Lan JY, Skeberdis VA, Jover T, Zheng X, Bennett MV, et al. 2001b. Activation of metabotropic glutamate receptor 1 accelerates NMDA receptor trafficking. J. Neurosci 21: 6058-6068. Lau LF, Huganir RL. 1995. Differential tyrosine phosphorylation of N‐methyl‐D‐aspartate receptor subunits. J Biol Chem 270: 20036-20041. Lee HK, Barbarosie M, Kameyama K, Bear MF, Huganir RL. 2000. Regulation of distinct AMPA receptor phosphorylation sites during bidirectional synaptic plasticity. Nature 405: 955-959. Lee SH, Liu L, Wang YT, Sheng M. 2002. Clatrin adaptor AP2 and NSF interact with overlapping sites of GluR2 and plays distinct roles in AMPA receptor trafficking and hippocampal LTD. Neuron 36: 661-674. Lei S, Czerwinska E, Czerwinski W, Walsh MP, McDonald JF. 2001. Regulation of NMDA receptor activity by F‐actin and myosin light chain kinase. J Neurosci 21: 8464-8472. Leonard AS, Hell JW. 1997. Cyclic AMP‐dependent protein kinase and protein kinase C phosphorylate N‐methyl‐D‐ aspartate receptors at different sites. J Biol Chem 272: 12107-12115. Lerma J. 2003. Roles and rules of kainate receptors in synaptic transmission. Nat Rev Neurosci 4: 481-495. Li B, Chen N, Luo T, Otsu Y, Murphy TH, et al. 2002. Differential regulation of synaptic and extra‐synaptic NMDA receptors. Nat Neurosci 5: 833-834. Li B, Otsu Y, Murphy TH, Raymond LA. 2003. Developmental decrease in NMDA receptor desensitization associated with shift to synapse and interaction with postsynaptic density‐ 95. J Neurosci 23: 11244-11254. Liao GY, Wagner DA, Hsu MH, Leonard JP. 2001. Evidence for direct protein kinase‐C mediated modulation of N‐methyl‐D‐aspartate receptor current. Mol Pharmacol 59: 960-964. Lisman J. 1989. A mechanism for the Hebb and the anti‐Hebb processes underlying learning and memory. Proc Natl Acad Sci USA 86: 9574-9578. Liu S‐QJ, Cul‐Candy SG. 2000. Synaptic activity at calcium‐ permeable AMPA receptors induces a switch in receptor subtype. Nature 405: 454-458. Lopez‐Bendito G, Shigemoto R, Fairen A, Lujan R. 2002. Differential distribution of group I metabotropic glutamate receptors during rat cortical development. Cereb Cortex 12: 625-638. Lozovaya NA, Grebenyuk SE, Tsintsadze TS, Feng B, Monaghan DT, et al. 2004. Extrasynaptic NR2B and NR2D subunits of NMDA receptors shape ‘superslow’ afterburst EPSC in rat hippocampus. J Physiol 558: 451-463.

437

438

17

Molecular organization and regulation of glutamate receptors in developing and adult mammalian CNS

Lu W, Man H, Ju W, Trimble WS, Mac Donald JF, et al. 2001. Activation of synaptic NMDA receptors induces membrane insertion of new AMPA receptors and LTP in cultured hippocampal neurons. Neuron 29: 243-254. Lujan R, Roberts JDB, Shigemoto R, Ohishi H, Somogyi P, 1997. Differential plasma membrane distribution of metabotropic glutamate receptors mGluR1a, mGluR2 and mGluR5, relative to neurotransmitter release sites. J Chem Neuroanat 13: 219-241. Luscher C, Xia H, Beattie EC, Carroll RC, von Zastrow M, et al. 1999. Role of AMPA receptor cycling in synaptic transmission and plasticity. Neuron 24: 649-658. Luyt K, Va´radi A, Molna´r E. 2003. Functional metabotropic glutamate receptors are expressed in oligodendrocyte progenitor cells. J Neurochem 84: 1452-1464. Luyt K, Va´radi A, Durant CF, Molna´r E. 2006. Oligodendroglial metabotropic glutamate receptors are developmentally regulated and involved in the prevention of apoptosis. J Neurochem 99: 641-656. Mammen AL, Kemeyama K, Roche KW, Huganir RL. 1997. Phosphorylation of the alpha‐amino‐3‐hydroxy‐5‐ methylisoxazole‐4‐propionic acid receptor GluR1 subunit by calcium/calmodulin‐dependent kinase II. J Biol Chem 272: 32528-32533. Man HY, Wang Q, Lu WY, Ju W, Ahmadian G, et al. 2003. Activation of PI3‐kinase is required for AMPA receptor insertion during LTP of mEPSCs in cultured hippocampal neurons. Neuron 38: 611-624. Martin LJ, Furuta A, Blacktone CD. 1998. AMPA receptor protein in developing rat brain: Glutamate receptor‐1 expression and localization change at regional, cellular, and subcellular levels with maturation. Neuroscience 83: 917-928. Martin S, Henley JH. 2004. Activity-dependent endocytic sorting of kainate receptors to recycling or degradation pathways. EMBO J 23: 4749-4759. Massey PV, Johnson BE, Moult PR, Auberson YP, Brown MW, et al. 2004. Differential roles of NR2A and NR2B-containing NMDA receptors in cortical long-term potentiation and long-term depression. J Neurosci 24: 7821-7828. Masu M, Tanade Y, Tsuchida K, Shigemoto R, Nakanishi S. 1991. Sequence and expression of a metabotropic glutamate receptor. Nature 349: 760-765. Matsuda S, Mikawa S, Hirai H. 1999. Phosphorylation of serine‐880 in GluR2 by protein kinase C prevents its C terminus from binding with glutamate receptor‐interacting protein. J Neurochem 73: 1765-1768. Mayer ML. 2005. Glutamate receptor ion channels. Curr Opin Neurobiol 15: 282-288. Mayer ML, 2006. Glutamate receptors at atomic resolution. Nature 440: 456-462.

Mayer ML, Armstrong N. 2004. Structure and function of glutamate receptor ion channels. Annu Rev Physiol 66: 161-181. Mayer ML, Westbrook GL, Guthrie PB. 1984. Voltage‐ dependent block by Mg2þ of NMDA responses in spinal cord neurons. Nature 309: 261-263. McDonald BJ, Chung HJ, Huganir RL. 2001. Identification of protein kinase C phosphorylation sites within the AMPA receptor GluR2 subunit. Neuropharmacology 41: 672-679. McIlhinney RAJ, Le Bourdelle`s B, Molna´r E, Tricaud N, Streit P, et al. 1998. Assembly intracellular targeting and cell surface expression of the human N‐methyl‐D‐aspartate receptor subunits NR1a and NR2A in transfected cells. Neuropharmacology 37: 1355-1367. McIlhinney RAJ, Molna´r E, Atack JR, Whiting PJ. 1996. Cell surface expression of the human N‐ethyl‐D‐aspartate receptor subunit 1a requires the co‐expression of the NR2A subunit in transfected cells. Neuroscience 70: 989-997. Minakami R, Iida K, Hirakawa N, Sugiyama H. 1995. The expression of two splice variants of metabotropic glutamate receptor subtype 5 in the rat brain and neuronal cells during development. J Neurochem 65: 1536-1542. Molna´r E, Baude A, Richmond SA, Patel PB, Somogyi P, et al. 1993. Biochemical and immunocytochemical characterization of antipeptide antibodies to a cloned GluR1 glutamate receptor subunit: Cellular and subcellular distribution in the rat forebrain. Neuroscience 53: 307-326. Molna´r E, McIlhinney RAJ, Baude A, Nusser Z, Somogyi P. 1994. Membrane topology of the GluR1 glutamate receptor subunit: Epitope mapping by site‐directed anti‐peptide antibodies. J Neurochem 63: 683-693. Molna´r E, Varadi A, McIlhinney RAJ, Ashcroft SJH. 1995. Identification of functional ionotropic glutamate receptor proteins in pancreatic b‐cells and islets of Langerhans. FEBS Lett 371: 253-257. Monyer H, Burnashev N, Laurie DJ, Sakmann B, Seeburg PH. 1994. Developmental and regional expression in the rat brain and functional properties of four NMDA receptors. Neuron 12: 529-540. Moon IS, Apperson ML, Kennedy MB, 1994. The major tyrosine‐phosphorylated protein in the postsynaptic density fraction is N‐methyl‐D‐aspartate receptor subunit 2B. Proc Natl Acad Sci USA 91: 3954-3958. Moriyoshi K, Masu M, Ishii T, Shigemoto R, Mizuno N, et al. 1991. Molecular cloning and characterization of the rat NMDA receptor. Nature 354: 31-37. Moult PR, Gladding CM, Sanderson TM, Fitzjohn SM, Bashir ZI, et al. 2006. Tyrosine phosphatases regulate AMPA receptor trafficking during metabotropic glutamate receptor‐mediated long‐term depression. J Neurosci 26: 2544-2554.

Molecular organization and regulation of glutamate receptors in developing and adult mammalian CNS Nakagawa T, Cheng Y, Ramm E, Sheng M, Walz T. 2005. Structure and different conformational states of native AMPA receptor complexes. Nature 433: 545-549. Nakajima Y, Yamamoto T, Nakayama T, Nakanishi S. 1999. A relationship between protein kinase C phosphorylation and calmodulin binding to the metabotropic glutamate receptor subtype 7. J Biol Chem 274: 27573-27577. Nakazawa T, Komai S, Tezuka T, Hisatsune C, Umemori H, et al. 2001. Characterization of Fyn‐mediated tyrosine phosphorylation sites on GluRepsilon 2 (NR2B) subunit of the N‐methyl‐D‐aspartate receptor. J Biol Chem 276: 693-699. Nasu‐Nishimura Y, Hurtado D, Braud S, Tze‐Tsang T, Isaac JTR, et al. 2006. Identification of an endoplasmic reticulum‐retention motif in an intracellular loop of the kainate receptor subunit KA2. J Neurosci 26: 7014-7021. Nicoletti F, Meek JL, Iadarola MJ, Chuang DM, Roth BL, et al. 1986. Coupling of inositol phospholipids metabolism with excitatory amino acid recognition sites in rat hippocampus. J Neurochem 46: 40-46. Nicoll RA, Tomita S, Bredt DS. 2006. Auxiliary subunits assist AMPA‐type glutamate receptors. Science 311: 1253-1256. Nishimune A, Isaac JTR, Molna´r E, Noe¨l J, Nash SR, et al. 1998. NSF binding to GluR2 regulates synaptic transmission. Neuron 21: 87-97. Noe¨l J, Ralph GS, Pickard L, Williams J, Molna´r E, et al. 1999. Surface expression of AMPA receptors in hippocampal neurons is regulated by an NSF‐dependent mechanism. Neuron 23: 365-376. Nong Y, Huang YQ, Ju W, Kalia LV, Ahmadian G, et al. 2003. Glycine binding primes NMDA receptor internalization. Nature 422: 302-307. Nowak L, Bregestovski P, Ascher P, Herbet A, Prochiantz A. 1984. Magnesium gated glutamate‐activated channels in mouse central neurons. Nature 307: 462-465. Nusser Z, Lujan R, Laude G, Roberts JDB, Molna´r E, et al. 1998. Cell type and pathway dependence of synaptic AMPA receptor number and variability in the hippocampus. Neuron 21: 545-559. O’Connor V, El Far O, Bofill‐Cardona E, Nanoff C, Freissmuth M, et al. 1999. Calmodulin dependence of presynaptic metabotropic glutamate receptor signalling. Science 286: 1180-1184. Oh MC, Derkach VA, Guire ES, Soderling TS. 2006. Extrasynaptic membrane trafficking regulated by GluR1 serine 845 phosphorylation primes AMPA receptors for long‐term potentiation. J Biol Chem 281: 752-758. Omkumar RV, Kiely MJ, Rosenstein AJ, Min KT, Kennedy MB. 1996. Identification of a phosphorylation site for calcium/calmodulin dependent protein kinase II in the NR2B subunit of the N‐methyl‐D‐aspartate receptor. J Biol Chem 271: 31670-31678.

17

Osten P, Khatri L, Perez JL, Kohr G, Giese G, et al. 2000. Mutagenesis reveals a role for ABP/GRIP binding to GluR2 in synaptic surface accumulation of the AMPA receptor, Neuron 27: 313-325. Osten P, Srivastava S, Inman GJ, Vilim FS, Khatri L, et al. 1998. The AMPA receptor GluR2 C terminus can mediate a reversible, ATP‐dependent interaction with NSF and alpha‐ and beta‐SNAPs, Neuron 21: 99-110. Passafaro M, Piech V, Sheng M. 2001. Subunit‐specific temporal and spatial patterns of AMPA receptor exocytosis in hippocampal neurons. Nat Neurosci 4: 917-926. Pe´rez‐Otan˜o I, Lujan R, Tavalin ST, Plomann M, Modregger J, et al. 2006. Endocytosis and synaptic removal of NR3A‐ containing NMDA receptors by PACSIN1/synapsin1. Nat Neurosci 9: 611-621. Petralia RS, Esteban JA, Wang YX, Partridge JG, Zhao HM, et al. 1999. Selective acquisition of AMPA receptors over postnatal development suggests a molecular basis for silent synapses. Nat Neurosci 2: 31-36. Petralia RS, Wang YX, Wenthold RJ. 1994a. Histological and ultrastructural localization of the kainate receptor subunits, KA2 and GluR6/7, in the rat nervous system using selective antipeptide antibodies. J Comp Neurol 349: 85-110. Petralia RS, Wang YX, Wenthold RJ. 1994c. The NMDA receptor subunits NR2A and NR2B show histological and ultrastructural localization patterns similar to those of NR1. J Neurosci 14: 6102-6120. Petralia RS, Yokotani N, Wenthold RJ. 1994b. Light and electron microscope distribution of the NMDA receptor subunit NMDAR1 in the rat nervous system using a selective anti‐peptide antibody. J Neurosci 14: 667-696. Pickard L, Noe¨l J, Duckworth JK, Fitzjohn SM, Henley JM, et al. 2001. Transient synaptic activation of NMDA receptors leads to the insertion of native AMPA receptors into hippocampal neuronal plasma membrane. Neuropharmacology 41: 700-713. Pickard L, Noe¨l J, Henley JM, Collingridge GL, Molna´r E. 2000. Developmental changes in synaptic AMPA and NMDA receptor distribution and AMPA receptor subunit composition in living hippocampal neurons. J Neurosci 20: 7922-7931. Plant K, Pelkey KA, Bortolotto ZA, Morita D, Terashima A, et al. 2006. Transient incorporation of native GluR2‐lacking AMPA receptors during hippocampal long‐term potentiation. Nat Neurosci 9: 602-604. Quinlan EM, Philpot BD, Huganir RL, Bear MF. 1999. Rapid, experience‐dependent expression of synaptic NMDA receptors in visual cortex in vivo. Nat Neurosci 2: 352-357. Racca C, Stephenson FA, Streit P, Roberts JDB, Somogyi P. 2000. NMDA receptor content of synapses in stratum radiatum of the hippocampal CA1 area. J Neurosci 20: 2512-2522.

439

440

17

Molecular organization and regulation of glutamate receptors in developing and adult mammalian CNS

Raymond LA, Blackstone CD, Huganir RL. 1993. Phosphorylation and modulation of recombinant GluR6 glutamate receptors by cAMP‐dependent protein kinase. Nature 361: 637-641. Ren Z, Riley NJ, Garcia EP, Sanders JM, Swanson GT, et al. 2003. Multiple trafficking signals regulate kainate receptor KA2 subunit surface expression. J Neurosi 23: 6608-6616. Roche KW, O’Brien RJ, Mammen AL, Bernhardt J, Huganir RL. 1996. Characterization of multiple phosphorylation sites on the AMPA receptor GluR1 subunit. Neuron 16: 1179-1188. Roche KW, Standley S, McCallum J, Dune Ly C, Ehlers MD, et al. 2001. Molecular determinants of NMDA receptor internalization. Nat Neurosci 4: 794-802. Roche KW, Tu JC, Petralia RS, Xiao B, Wenthold RJ, et al. 1999. Homer 1b regulates the trafficking of group I metabotropic glutamate receptors. J Biol Chem 274: 25953-25957. Romano C, Yang WL, O’Malley KL. 1996a. Metabotropic glutamate receptor 5 is a disulfide‐linked dimmer. J Biol Chem 271: 28612-28616. Romano C, van den Pol AN, O’Malley KL. 1996b, Enhanced early developmental expression of the metabotropic glutamate receptor mGluR5 in rat brain: Protein, mRNA splice variants, and regional distribution. J Comp Neurol 367: 403-412. Romano C, Smout S, Miller JK, O’Malley KL. 2002. Developmental regulation of metabotropic glutamate receptor 5b protein in rodent brain. Neuroscience 111: 693-698. Ruano D, Lambolez B, Rossier J, Paternain AV, Lerma J. 1995. Kainate receptor subunits expressed in single cultured hippocampal neurones: Molecular and functional variants by RNA editing. Neuron 14: 1009-1017. Ruiz A, Sachidhanandam S, Utvik JK, Coussen F, Mulle C. 2005. Distinct subunits in heteromeric kainate receptors mediate ionotropic and metabotropic function at hippocampal mossy fiber synapses. J Neurosci 25: 11710-11718. Sa´nchez‐Pe´rez AM, Felipo V. 2005. Serines 890 and 896 of the NMDA receptor subunit NR1 are differentially phosphorylated by protein kinase C isoforms. Neurochem Int 47: 84-91. Sasaki YF, Rothe T, Premkumar LS, Das S, Cui J, et al. 2002. Characterization and composition of the NR3A subunit of the NMDA receptor in recombinant systems and primary cortical neurons. J Neurophysiol 87: 2052-2063. Schoepp DD, Jane DE, Monn JA. 1999. Pharmacological agents acting at subtypes of metabotropic glutamate receptors. Neuropharmacology 38: 1431-1476. Scott DB, Blanpied TA, Ehlers MD. 2003. Coordinated PKA and PKC phosphorylation suppresses RXR‐mediated ER retention and regulates the surface delivery of NMDA receptors. Neuropharmacology 45: 755-767.

Seidenman KJ, Steinberg JP, Huganir R, Malinow R. 2003. Glutamate receptor subunit 2 serine 880 phosphorylation modulates synaptic transmission and mediates plasticity in CA1 pyramidal cells. J Neurosci 23: 9220-9228. Serge A, Fourgeaud L, Hemar A, Choquet D. 2002. Receptor activation and homer differentially control the lateral mobility of metabotropic glutamate receptor 5 in the neuronal membrane. J Neurosci 22: 3910-3920. Shen L, Liang F, Walensky LD, Huganir RL. 2000. Regulation of AMPA receptor GluR1 subunit surface expression by a 4.1N‐linked actin cytoskeletal association. J Neurosci 20: 7932-7940. Shi S, Hayashi Y, Esteban JA, Malinow R. 2001. Subunit‐ specific rules governing AMPA receptor trafficking to synapses in hippocampal pyramidal neurons. Cell 105: 331-343. Shigemoto R, Kinoshita A, Wada E, Nomura S, Ohishi H, et al. 1997. Differential presynaptic localization of metabotropic glutamate receptor subtypes in the rat hippocampus. J Neurosci 17: 7503-7522. Shigemoto R, Kulik A, Roberts JD, Ohishi H, Nusser Z, et al. 1996. Target‐cell‐specific concentration of a metabotropic glutamate receptor in the presynaptic active zone Nature 381: 523-525. Shigemoto R, Nakanishi S, Mizuno N. 1992. Distribution of the mRNA for a metabotropic glutamate receptor (mGluR1) in the central nervous system: An in situ hybridization study in adult and developing rat. J Comp Neurol 322: 121-135. Skeberdis VA, Chevaleyre V, Lau CG, Goldberg JH, Pettit DL, et al. 2006. Protein kinase A regulates calcium permeability of NMDA receptors. Nat Neurosci 9: 501-510. Skeberdis VA, Lan J, Zheng X, Zukin RS, Bennett MV. 2001. Insulin promotes rapid delivery of N‐methyl‐D‐aspartate receptors to the cell surface by exocytosis. Proc Natl Acad Sci USA 98: 3561-3566. Sladeczek F, Pin JP, Recasens M, Bockaert J, Weiss S. 1985. Glutamate stimulates inositol phosphate formation in striatal neurones. Nature 317: 717-719. Smith TC, Wang LY, Howe JR. 1999. Distinct kainate receptor phenotypes in immature and mature mouse cerebellar granule cells. J. Physiol 517: 51-58. Snyder EM, Philpot BD, Huber KM, Dong X, Fallon JR, et al. 2001. Internalization of ionotropic glutamate receptors in response to mGluR activation. Nat Neurosci 4: 1079-1085. Standley S, Roche KW, McCallum J, Sans N, Wenthold RJ. 2000. PDZ domain suppression of an ER retention signal in NMDA receptor NR1 splice variants. Neuron 28: 887-898. Steinberg JP, Takamiya K, Shen Y, Xia J, Rudio ME, et al. 2006. Targeted in vivo mutations of the AMPA receptor subunit

Molecular organization and regulation of glutamate receptors in developing and adult mammalian CNS GluR2 and its interacting protein PICK1 eliminate cerebellar long‐term depression. Neuron 49: 845-860. Stocca G, Vicini S. 1998. Increased contribution of NR2A subunit to synaptic NMDA receptors in developing rat cortical neurons J Physiol 507: 13-24. Sun L, Margolis FL, Shipley MT, Lidow MS. 1998. Identification of a long variant of mRNA encoding the NR3 subunit of the NMDA receptor: Its regional distribution and developmental expression in the rat brain. FEBS Lett 441: 392-396. Sun X, Zhao Y, Wolf ME. 2005. Dopamine receptor stimulation modulates AMPA receptor synaptic insertion in prefrontal cortex neurons. J Neurosci 25: 7342-7351. Takumi Y, Ramı´rez‐Leo´n V, Laake P, Rinvik E, Ottersen OP. 1999. Different modes of expression of AMPA and NMDA receptors in hippocampal synapses. Nat Neurosci 2: 618-624. Tamaru Y, Nomura S, Mizuno N, Shigemoto R. 2001. Distribution of metabotropic glutamate receptor mGluR3 in the mouse CNS: Differential location relative to pre‐ and postsynaptic sites. Neuroscience 106: 481-503. Tanaka J, Matsuzaki M, Tarusawa E, Momiyama A, Molna´r E, et al. 2005. Number and density of AMPA receptors in single synapses in immature cerebellum. J Neurosci 25: 799-807. Tang YP, Shimizu E, Dube GR, Rampon C, Kerchner GA, et al. 1999. Genetic enhancement of learning and memory in mice. Nature 401: 63-69. Tarnawa I, Vizi ES. 1998. 2,3‐Benzodiazepine AMPA antagonists. Restor Neurol Neurosci 13: 41-57. Terashima A, Cotton L, Dev KK, Meyer G, Zaman S, et al. 2004. Regulation of synaptic strength and AMPA receptor subunit composition by PICK1. J Neurosci 24: 5381-5390. Tigaret CM, Thalhammer A, Rast GF, Specht CG, Auberson YP, et al. 2006. Subunit dependencies of N‐methyl‐D‐ aspartate (NMDA) receptor‐induced a‐amino‐3‐hydroxy‐ 5‐methyl‐4‐isoxazolepropionic acid (AMPA) receptor internalization. Mol Pharmacol 69: 1251-1259. Tingley WG, Ehlers MD, Kameyama K, Doherty C, Ptak JB, et al. 1997. Characterization of protein kinase A and protein kinase C phosphorylation of the N‐methyl‐D‐aspartate receptor NR1 subunit using phosphorylation site‐specific antibodies. J Biol Chem 272: 5157-5166. To¨nnes J, Stierli B, Cerletti C, Behrmann JT, Molna´r E, et al. 1999. Regional distribution and developmental changes of GluR1‐flop protein revealed by monoclonal antibody in rat brain. J Neurochem 73: 2195-2205. Tovar KR, Westbrook GL. 2002. Mobile NMDA receptors at hippocampal synapses. Neuron 34: 255-264.

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Triller A, Choquet D. 2005. Surface trafficking of receptors between synaptic and extrasynaptic membranes: And yet they do move! Trends Neurosci 28: 133-139. Tsuchiya D, Kunishima N, Kamiya N, Jingami H, Morikawa K. 2002. Structural views of the ligand‐binding cores of a metabotropic glutamate receptor complexed with an antagonist and both glutamate and Gb3þ. Proc Natl Acad Sci USA 99: 2660-2665. Vissel B, Krupp JJ, Heinemann SF, Westbrook GL. 2001. A use‐dependent tyrosine dephosphorylation of NMDA receptors is independent of ion flux. Nat Neurosci 4: 587-596. Vizi ES. 2000. Role of high‐affinity receptors and membrane transporters in nonsynaptic communication and drug action in the central nervous system. Pharmacol Rev 52: 63-89. Vizi ES, Mike A´. 2006. Nonsynaptic receptors for GABA and Glu. Curr Top Med Chem 6: 941-948. Vizi ES, Mike A, Tarnawa I. 1996. 2,3 Benzodiazepines (GKI 52466 and analogs): Negative allosteric modulators of AMPA receptors. CNS Drug Rev 2: 91-126. Wang LY, Taverna FA, Huang XP, Mac Donald JF, Hampson DR. 1993. Phosphorylation and modulation of a kainate receptor (GluR6) by cAMP‐dependent protein kinase. Science 259: 1173-1175. Washbourne P, Bennett JE, McAllister AK. 2002. Rapid recruitment of NMDA receptor transport packets to nascent synapses. Nat Neurosci 5: 751-759. Watanabe M. 1997. Developmental dynamics of gene expression for NMDA receptor channel. The Ionotropic Glutamate Receptors. Monaghan DT, Wenthold RJ, editors. Humana Press, New Jersey. Watkins JC, Jane DE. 2006. The glutamate story. Br J Pharmacol 147: S100-S108. Wenthold RJ, Prybylowski K, Standley S, Sans N, Petralia RS. 2003. Trafficking of NMDA receptors. Annu Rev Pharmacol Toxicol 43: 335-358. Winder DG, Conn PJ. 1996. Roles of metabotropic glutamate receptors in glial function and glial‐neuronal communication. J Neurosci Res 46: 131-137. Wu H, Nash JE, Zamarano P, Garner CC. 2002. Interaction of SAP97 with minus‐end‐directed actin motor myosin VI. Implications for AMPA receptor trafficking. J Biol Chem 277: 30928-30934. Xiao MY, Zhou Q, Nicoll RA. 2001. Metabotropic glutamate receptor activation causes a rapid redistribution of AMPA receptors. Neuropharmacology 41: 664-671. Yang M, Leonard JP. 2001. Identification of mouse NMDA receptor subunit NR2A C‐terminal tyrosine sites phosphorylated by coexpression with v‐Src. J Neurochem 77: 580-588.

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Sympathetic and Peptidergic Innervation: Major Role at the Neural–Immune Interface

I. J. Elenkov . A. Tagliani

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The Neural–Immune Interface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 444

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Lymphocyte Traffic and Proliferation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 445

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NK‐Cell Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 446

4 4.1 4.2 4.3 4.4

Pro‐ and Anti‐Inflammatory and Th1‐ and Th2‐Type Cytokines . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 446 NE and Epinephrine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 446 Adenosine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 447 ATP and NPY . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 447 SP and CGRP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 448

5 Systemic versus Local Effects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 448 5.1 CRH/SP–Mast Cell–Histamine Axis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 448 6

Antibody Production . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 450

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Conclusion and Clinical Implications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 450

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Sympathetic and peptidergic innervation: Major role at the neural–immune interface

Abstract: Lymphoid organs receive extensive sympathetic and peptidergic/sensory innervation. Neurotransmitters and neuropeptides, released by the sympathetic and peptidergic nerve fibers in the vicinity of immunocompetent cells, and circulating epinephrine, secreted by the adrenal medulla affect major immune functions such as secretion of cytokines and antibodies, selection of T‐helper (Th)1 or Th2 responses, lymphocyte activity, proliferation and traffic. Catecholamines (CAs), ATP, and adenosine inhibit the production of interleukin (IL‐12), tumor necrosis factor (TNF)‐a, and interferon (IFN)‐g, whereas they stimulate the production of IL‐10. Thus, systemically, sympathetic neurotransmitters may induce a Th2 shift, and neuropeptide Y may further amplify this effect. In certain local responses, however, CAs induce IL‐1, IL‐6, IL‐8, and TNF‐a production; substance P upregulates IL‐1, IL‐12, and TNF‐a production, whereas ATP via the P2X7 receptor may activate the posttranslational processing of IL‐1b and IL‐18. This might be aimed to localize the inflammatory response through induction of neutrophil and monocyte accumulation and stimulation of macrophage activity. Systemically, however, the activation of the sympathetic nervous system during an immune response may suppress Th1 responses, and, hence protect the organism from the detrimental effects of proinflammatory cytokines and other products of activated macrophages. Thus, a dysfunctional neural–immune interface may play a role in the pathogenesis of various common human immune‐related diseases. List of Abbreviations: ADO, adenosine; APC, antigen‐presenting cells; BALT, Bronchus‐associated lymphoid tissue; BCDF, B‐cell differentiation factor; CGRP, calcitonin gene‐related peptide; CRH, corticotropin‐releasing hormone; CRP, C‐reactive protein; GALT, Gut‐associated lymphoid tissue; PHI, peptide histidine isoleucine; SLE, systemic lupus erythematosus; SNS, sympathetic nervous system; TGF, transforming growth factor; VIP, vasoactive intestinal polypeptide

1

The Neural–Immune Interface

The brain affects the immune system through the neuroendocrine humoral outflow via the pituitary, and through direct neuronal influences via the sympathetic, peptidergic/sensory, and parasympathetic (cholinergic) innervation of peripheral tissues. Lymphoid organs, and particularly their parenchyma, similar to smooth muscles of the vasculature, receive predominantly sympathetic and peptidergic/sensory innervation. Generally, zones of T cells, macrophages, and plasma cells are richly innervated, whereas nodular and follicular zones of developing or mature B cells are poorly innervated (Felten et al., 1985). In lymphoid organs, norepinephrine (NE), the major sympathetic neurotransmitter is released nonsynaptically, that is, from varicose axon terminals, which do not make synaptic contacts – NE released from perivascular or connective tissue septa plexuses of nerve terminals diffuse away through surrounding adventitia or collagenous fibrils in a paracrine fashion. Thus, NE may play a modulatory role in signal transmission at the sympathetic–immune interface (Vizi et al., 1995; Elenkov et al., 2000b). Sympathetic/neuropeptide Y (NPY)‐positive fibers predominantly supply the vasculature, where they mainly occur as perivascular plexuses, and both NE and NPY released from these fibers control blood flow and lymphocyte traffic. They branch off only rarely to run into the lymphoid parenchyma (Felten et al., 1985). NPY is coreleased with NE on sympathetic nervous system (SNS) activation – particularly in conditions of high sympathetic activity large dense‐cored vesicles release both NPY and NE (Lundberg et al., 1989). NPY does not usually act as a genuine cotransmitter but rather as a prejunctional or postjunctional modulator of the release or the effects of the principal transmitters, NE and ATP. In many tissues, the major action of NPY is to enhance the postjunctional response of NE and ATP. In many tissues, ATP is costored with NE and NPY in the sympathetic nerve terminals (Burnstock, 1997). The sympathetic nerves most likely release ATP transiently, only at the beginning of a train of nerve stimulation. The release of NE occurs later in the train and, once started, is maintained throughout the course of nerve stimulation (Westfall et al., 2002). In blood vessels, ATP is particularly abundant, and the proportion of ATP to NE is extremely variable in different blood vessels beds. Once released, ATP is rapidly breakdown to adenosine (ADO) by extracellular nucleotidases (Burnstock, 1997; Westfall et al., 2002). Ischemic‐like condition releases both NE and ATP in the rat spleen (Sperlagh et al., 2000); further

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studies, however, are needed to clarify the release of ATP and its source in lymphoid organs under more physiologic conditions. In lymphoid organs, the peptidergic/sensory innervation is confined mostly to the parenchyma (Weihe et al., 1991). The most abundant peptides are tachykinins (substance P, neurokinin A), calcitonin gene‐ related peptide (CGRP), and vasoactive intestinal polypeptide/peptide histidine isoleucine (VIP/PHI). Double immunofluorescence reveals coexistence of tachykinins with CGRP and of TH and NPY. The coexistence of peptides with peptides and of NPY with markers of the catecholamine pathway conforms to the general scheme described for the peripheral innervation of other organs. Similar to other organs, the tachykinins/CGRP fibers most likely have sensory origins. As a general pattern, and as in the case of noradrenergic innervation, a close spatial relationship between peptidergic nerve fibers and mast cells, T cells and macrophages is observed (Weihe et al., 1991). Peptidergic nerves also appear to be sparse in pure B‐cell regions. Neuro–mast cell contacts are often relatively seen in all lymphoid organs with the exception of the spleen (Weihe et al., 1991). Gut‐associated lymphoid tissue (GALT) and bronchus‐associated lymphoid tissue (BALT) also receive extensive sympathetic and peptidergic innervation (Felten et al., 1985). The most copious peptides are SP and CGRP closely overlapping anatomically, but not necessarily colocalized in the sensory innervation, and VIP, present in the cholinergic innervation. Neuropeptides are found in very large amounts in these tissues, particularly SP, VIP, and somatostatin. Although this issue is not extensively studied, the available information suggests that in GALT including Peyer’s patches that represent clusters of lymphoid nodules in the intestines, and the appendix of the rabbit, varicose noradrenergic fibers arborize profusely in the interdomal region of the lamina propria. Here, fibers follow small vessels and branch freely in the parenchyma among fields of lymphoid cells, usually not in association with blood vessels (Bellinger et al., 1992). Nerves predominate in T‐cell zones of lymphoid aggregates, where they contain neuropeptides and the sympathetic neurotransmitter NE. As in other lymphoid tissues, intestinal mucosal mast cells lying immediately under the epithelium are apparently selectively associated with enteric nerves. In these tissues, the noradrenergic varicosities are also adjacent to serotonergic enterochromaffin cells (Felten et al., 1985).

2

Lymphocyte Traffic and Proliferation

Administration of catecholamines (CAs, NE, and epinephrine) in humans induces a quick ( Figure 18-1). Furthermore, CAs (through b2/b3‐ARs) upregulate IL‐6 production by human adipocytes (Kaplanski et al., 1993; Engstad et al., 1999; Mohamed‐Ali et al., 2001; Vicennati et al., 2002). IL‐6 is the major inducer of C‐reactive protein (CRP) production by the liver and CAs enhance this induction to a greater or lesser extent (Baumann and Gauldie, 1994). Interestingly, chronic b‐AR stimulation induces myocardial, but not systemic, elaboration of TNF‐a, IL‐1b, and IL‐6 (Murray et al., 2000; Li et al., 2001). Several studies suggest that CAs suppress the tumoricidal and the cytotoxic state of activated macrophages, mostly through inhibition of TNF‐a and IL‐1, and potentiation of the production of IL‐10 (Koff and Dunegan, 1985, 1986). In contrast, both NE and epinephrine were reported to stimulate murine resident peritoneal macrophages to suppress the growth of Mycobacterium avium, an effect mediated by a2‐ARs (Miles et al., 1996). The apparent discrepancy between stimulatory and inhibitory activities of CAs may be attributed to the state of activation of macrophage populations: antigen challenge and activation of macrophages may result in an increase in b‐receptors and suppression of the response. It is highly likely, however, that there is a transient stage of differentiation, when monocytes (during maturation to macrophages) lose their b‐AR responsiveness (see Baker and Fuller, 1995). Thus, naı¨ve cells may preferentially express a‐ARs, which will result in stimulation of macrophage activity. In fact, the a2‐ARs‐mediated stimulatory effect of NE on TNF‐a production was observed in peritoneal macrophages elicited from specific pathogen‐free mice (Spengler et al., 1990), a condition that may reflect naı¨ve, antigen‐inexperienced macrophages.

5.1 CRH/SP–Mast Cell–Histamine Axis Peripherally produced corticotropin‐releasing hormone (CRH) acts as a local autocrine and paracrine proinflammatory agent (Karalis et al., 1991). Immunoreactive CRH is identified locally in experimental carrageenin‐ induced subcutaneous aseptic inflammation, streptococcal cell wall‐ and adjuvant‐induced arthritis, and in

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. Figure 18-1 Simplified scheme of the complex interactions between CAs, neuropeptides, and the CRH/SP–mast cell–histamine axis, and their pro- and anti-inflammatory effects in certain local responses (see text; from Elenkov, 2003). Solid lines represent stimulation, whereas dashed lines inhibition. Abbreviations: CGRP, calcitonin gene-related peptide; CRH, corticotropin-releasing hormone (peripheral); EPI, epinephrine; IL, interleukin; NE, norepinephrine; SP, substance P; TNF, tumor necrosis factor

human tissues from patients with rheumatoid arthritis (RA), autoimmune thyroid disease, and ulcerative colitis. CRH in early inflammation is of peripheral, mostly peptidergic/sensory nerve rather than immune‐cell origin. Recent evidence also indicates that urocortin, a new member of the CRH family that acts through the family of CRH receptors, is overexpressed in the synovia of patients with RA and this is related to inflammatory activity (Karalis et al., 1991; Crofford et al., 1993; Elenkov et al., 1999; Kohno et al., 2001). Peripheral CRH has vascular permeability enhancing and vasodilatory actions. CRH administration causes major peripheral vasodilation manifested as flushing and increased blood flow and hypotension (Udelsman et al., 1986). An intradermal CRH injection induces a marked increase of vascular permeability and mast cell degranulation, mediated through CRH type 1 receptors (Theoharides et al., 1998). It appears that the mast cell is a major target of immune CRH. This concept has an anatomic prerequisite in blood vessels and the lymphoid parenchyma plexuses of nerve fibers (noradrenergic and peptidergic) are closely associated with clusters of mast cells (cf. Elenkov et al., 2000b). SP and peripheral CRH, which are released from sensory peptidergic neurons, are two of the most potent mast cell secretagogues (Foreman, 1987; Church et al., 1989; Theoharides et al., 1995, 1998). Peripheral CRH and SP activates mast cells via a CRH type 1 and NK1 receptor‐dependent mechanism, leading to release of histamine and other contents of the mast cell granules that subsequently may cause vasodilatation, increased vascular permeability, and other manifestations of inflammation. Thus, the activation of CRH/SP–mast cell–histamine axis through stimulation of H1 receptors may induce acute inflammation and allergic reactions, while through activation of H2 receptors it may induce suppression of Th1 responses and a Th2 shift. This adds further complexity to the local effects of hormones, neurotransmitters, and neuropeptides in conjunction with local mediators of inflammation (> Figure 18-1).

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Sympathetic and peptidergic innervation: Major role at the neural–immune interface

Antibody Production

When B cells and Th cells are exposed to Th cell‐dependent antigens, NE, through stimulation of b2 receptors, exerts an enhancing effect on B‐cell antibody (Ab) production (Sanders and Munson, 1985; Sanders et al., 1997). IFN‐g‐producing Th1 cells induce B cells to produce IgG2a (in humans, IgG1), whereas IL‐4‐producing Th2 cells induce B cells to produce IgE and IgG1 (in humans, IgG4) (Fearon and Locksley, 1996). Thus, the inhibition of IFN‐g production by Th1 cells induced by the b2‐AR agonist terbutaline is associated with subsequent suppressed IgG2a production by mouse B cells (Sanders et al., 1997). b‐AR agonists also potentiate IL‐4‐induced IgE production by human mononuclear cells, while they inhibit IFN‐g production by these cells (Coqueret et al., 1995). Thus, CAs may induce a switch to IgE, and to IgG1 (in rodents) or IgG4 (in humans) subclasses production. The enhancement of Ab production further supports the hypothesis that CAs mediate a Th2 shift that potentiates humoral immunity.

7

Conclusion and Clinical Implications

Neural–immune interactions are undoubtedly complex. Recent studies suggest that endogenous CAs modulate the function of primary lymphoid organs, such as the bone marrow and the thymus (Vizi et al., 1995; Maestroni, 1998a, b; Cavallotti et al., 2002). However, the role of the sympathetic innervation in regulation of hematopoiesis and thymocyte development remains poorly understood. In addition, there is almost complete lack of knowledge how CAs might affect mucosal immunity. Overall, there is substantial evidence indicating that local or systemic CAs, ATP, ADO, and NPY are involved in fine‐ tuning of immune responses. The effects of the sympathetic neurotransmitters/neuromodulators are quick within minutes. This modulation might be ideally designed for quick adjustment of lymphoid cell traffic, cytokine/chemokine production, or cellular responsiveness and activity. Although interest in the Th2 response was initially directed at its protective role in helminthic infections and its pathogenic role in allergy, this response may have important regulatory functions in countering the tissue‐damaging effects of macrophages and Th1 cells (Fearon and Locksley, 1996). Some proinflammatory cytokines, and particularly IL‐1, via its central effects stimulate the SNS output (Besedovsky et al., 1983; Dunn, 1988). Thus, an excessive immune response and the subsequent activation of the SNS, and the release of CAs in the periphery, may trigger a mechanism that inhibits, systemically, Th1 functions and proinflammatory cytokine production, but potentiates Th2 and anti‐inflammatory responses (Elenkov et al., 1996, 2000a, b; Elenkov and Chrousos, 1999). This appears to be complemented by locally released ATP, ADO, while NPY may amplify these effects. This mechanism may protect the organism from systemic ‘‘overshooting’’ with type 1/proinflammatory cytokines and other products of activated macrophages with tissue‐damaging potential. On the other hand, in certain local responses, and under certain conditions, CAs may actually boost regional innate immune responses in a transitory fashion, through induction of IL‐8, IL‐6, IL‐1, and TNF‐a production, and through short‐term increase of NK cell, monocyte, and neutrophil numbers. In addition, ATP via stimulation of the P2X7 receptor may activate the posttranslational processing and subsequent release of IL‐1b and IL‐18. This might be aimed to localize the inflammatory response via stimulation of neutrophils accumulation and stimulation of macrophage activity. Although a complete discussion is beyond the scope of this chapter, an abnormal sympathetic–immune interface or activity of the SNS may play a role in the pathogenesis of infections, autoimmune and atopic/allergic reactions, and tumor growth. Acute or chronic stress and subsequent CAs‐induced Th2 shift might specifically increase the susceptibility of the individual to intracellular infections, the defense against which is primarily through Th1‐ regulated cellular immunity – for example, mycobacterial, Helicobacter pylori, HIV, or common cold viral infections. The stress‐induced Th2 shift may also trigger herpes simplex viral reactivation (Cohen et al., 1991; Elenkov et al., 1996; Lerner, 1996; Padgett et al., 1998; Elenkov and Chrousos, 1999; Levenstein et al., 1999; Sainz et al., 2001). Additionally, NE directly accelerate HIV‐1 replication, while some of the instability in some inflammatory responses, such as leprosy, might be secondary to the damage of sensory

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C‐ and sympathetic nerve fibers and dysregulation of inflammation (Cole et al., 1998, 2001; Rook et al., 2002). Major injury (serious traumatic injury and major burns or major surgical procedures) often triggers a ‘‘sympathetic storm’’ – the subsequent massive release of CAs and ADO via an induction of a Th2 shift may contribute to the severe immunosuppression and the severe infectious complications observed in these conditions (O’Sullivan et al., 1995; Woiciechowsky et al., 1998; Elenkov and Chrousos, 1999; Elenkov et al., 2000a, b; Munford and Tracey, 2002). A hypoactive SNS system may facilitate or sustain the Th1 shift, observed in autoimmune diseases, such as RA or multiple sclerosis (MS). An additional factor might be the preponderance of about 10:1 for primary sensory, SP‐positive fibers as compared with sympathetic fibers in synovial tissues of RA patients (> Figure 18-2). Alternatively, SNS hyperactivity may intensify the Th2 shift and induce or facilitate flares of systemic lupus erythematosus (SLE) (Wilder, 1995; Elenkov and Chrousos, 1999; Elenkov et al., 2000a, b, 2001; Miller et al., 2000; Straub and Cutolo, 2001).

. Figure 18-2 Role of systemic and local neuroendocrine factors in the pathogenesis of rheumatoid arthritis. Patients with rheumatoid arthritis have hypoactive HPA axis and SNS in the settings of severe chronic inflammation, characterized by increased production of IL-1, IL-6, and TNF-a. The hypoactivity of the stress system may sustain and facilitate the Th1 shift observed in rheumatoid arthritis and further promote the local inflammation. Locally, the preponderance of primary sensory, SP-positive fibers as compared with sympathetic fibers and the overexpression of CRH and urocortin result in a dominance of autocrine and paracrine proinflammatory factors in the synovium of rheumatoid arthritis patients. Solid lines represent stimulation, whereas dashed lines inhibition. Abbreviations: APC, antigen-presenting cell; CRH, corticotropin-releasing hormone (peripheral); EPI, epinephrine; HPA, hypothalamic–pituitary–adrenal axis; IL, interleukin; NE, norepinephrine; SNS, sympathetic nervous system; SP, substance P; Th, T-helper lymphocyte; TNF, tumor necrosis factor

Allergic reactions of type 1 hypersensitivity (atopy), such as asthma, eczema, hay fever, urticaria, and food allergy, are characterized by dominant Th2 responses, overproduction of histamine, and a shift to IgE production. The effects of stress on atopic reactions are complex at multiple levels and can be in either direction. CAs acting at the level of APCs and lymphocytes may induce a Th2 shift, and thus facilitate or sustain atopic reactions; however, this can be antagonized by their effects on the mast cell (Barnes et al., 1980;

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Elenkov and Chrousos, 1999; Marshall and Agarwal, 2000). In addition, the activation of CRH/SP–mast cell–histamine axis may also contribute to inflammation in allergic reactions. Low levels of IL‐12 and local overproduction of IL‐10 and TGF‐b have been associated with tumor growth (Colombo et al., 1996; Chouaib et al., 1997). These data suggest that CAs‐ and ADO‐induced inhibition of IL‐12 and potentiation of IL‐10 and TGF‐b production, and subsequent suppression of cellular immunity may contribute to the increased growth of certain tumors (Hoskin et al., 1994; Blay et al., 1997; Li et al., 1997; Shakhar and Ben‐Eliyahu, 1998). Clearly, these hypotheses require further investigation, but the answers should provide critical insights into mechanisms underlying a variety of common human diseases.

References Baker AJ, Fuller RW. 1995. Loss of response to betaadrenoceptor agonists during the maturation of human monocytes to macrophages in vitro. J Leukoc Biol 57: 395-400. Barnes P, FitzGerald G, Brown M, Dollery C. 1980. Nocturnal asthma and changes in circulating epinephrine, histamine, and cortisol. N Engl J Med 303: 263-267. Bartik MM, Bauman GP, Brooks WH, Roszman TL. 1994. Costimulatory signals modulate the antiproliferative effects of agents that elevate cAMP in T cells. Cell Immunol 158: 116-130. Baumann H, Gauldie J. 1994. The acute phase response. Immunol Today 15: 74-80. Bedoui S, Kawamura N, Straub RH, Pabst R, Yamamura T, et al. 2003. Relevance of neuropeptide Y for the neuroimmune crosstalk. J Neuroimmunol 134: 1-11. Bellinger DL, Lorton D, Felten SY, Felten DL. 1992. Innervation of lymphoid organs and implications in development, aging, and autoimmunity. Int J Immunopharmacol 14: 329-344. Benschop RJ, Rodriguez-Feuerhahn M, Schedlowski M. 1996. Catecholamine-induced leukocytosis: Early observations, current research, and future directions. Brain Behav Immun 10: 77-91. Besedovsky H, Del Rey A, Sorkin E, Da Prada M, Burri R, et al. 1983. The immune response evokes changes in brain noradrenergic neurons. Science 221: 564-566. Blay J, White TD, Hoskin DW. 1997. The extracellular fluid of solid carcinomas contains immunosuppressive concentrations of adenosine. Cancer Res 57: 2602-2605. Borger P, Hoekstra Y, Esselink MT, Postma DS, Zaagsma J, et al. 1998. Beta-adrenoceptor-mediated inhibition of IFN-gamma, IL-3, and GM-CSF mRNA accumulation in activated human T lymphocytes is solely mediated by the beta2-adrenoceptor subtype. Am J Respir Cell Mol Biol 19: 400-407. Broug-Holub E, Persoons JH, Schornagel K, Mastbergen SC, Kraal G. 1998. Effects of stress on alveolar macrophages:

A role for the sympathetic nervous system. Am J Respir Cell Mol Biol 19: 842-848. Burnstock G. 1997. The past, present and future of purine nucleotides as signalling molecules. Neuropharmacology 36: 1127-1139. Cavallotti D, Artico M, Iannetti G, Cavallotti C. 2002. Occurrence of adrenergic nerve fibers in human thymus during immune response. Neurochem Int 40: 211-221. Chakfe Y, Seguin R, Antel JP, Morissette C, Malo D, et al. 2002. ADP and AMP induce interleukin-1beta release from microglial cells through activation of ATP-primed P2X7 receptor channels. J Neurosci 22: 3061-3069. Chambers DA, Cohen RL, Perlman RL. 1993. Neuroimmune modulation: Signal transduction and catecholamines. Neurochem Int 22: 95-110. Chouaib S, Asselin-Paturel C, Mami-Chouaib F, Caignard A, Blay JY. 1997. The host-tumor immune conflict: From immunosuppression to resistance and destruction. Immunol Today 18: 493-497. Church MK, Lowman MA, Robinson C, Holgate ST, Benyon RC. 1989. Interaction of neuropeptides with human mast cells. Int Arch Allergy Appl Immunol 88: 70-78. Cohen S, Tyrrell DA, Smith AP. 1991. Psychological stress and susceptibility to the common cold [see comments]. N Engl J Med 325: 606-612. Cole SW, Korin YD, Fahey JL, Zack JA. 1998. Norepinephrine accelerates HIV replication via protein kinase A-dependent effects on cytokine production. J Immunol 161: 610-616. Cole SW, Naliboff BD, Kemeny ME, Griswold MP, Fahey JL, et al. 2001. Impaired response to HAART in HIV-infected individuals with high autonomic nervous system activity. Proc Natl Acad Sci USA 98: 12695-12700. Colombo MP, Vagliani M, Spreafico F, Parenza M, Chiodoni C, et al. 1996. Amount of interleukin 12 available at the tumor site is critical for tumor regression. Cancer Res 56: 2531-2534. Coqueret O, Dugas B, Mencia-Huerta JM, Braquet P. 1995. Regulation of IgE production from human mononuclear

Sympathetic and peptidergic innervation: Major role at the neural–immune interface cells by beta 2-adrenoceptor agonists [see comments]. Clin Exp Allergy 25: 304-311. Coqueret O, Lagente V, Frere CP, Braquet P, Mencia-Huerta JM. 1994. Regulation of IgE production by beta 2-adrenoceptor agonists. Ann N Y Acad Sci 725: 44-49. Crofford LJ, Sano H, Karalis K, Friedman TC, Epps HR, et al. 1993. Corticotropin-releasing hormone in synovial fluids and tissues of patients with rheumatoid arthritis and osteoarthritis. J Immunol 151: 1587-1596. Dunn AJ. 1988. Systemic interleukin-1 administration stimulates hypothalamic norepinephrine metabolism parallelling the increased plasma corticosterone. Life Sci 43: 429-435. Elenkov IJ. January 7, 2003. http://www.endotext.org/adrenal/ adrenal28/adrenalframe28.htm Elenkov IJ, Chrousos GP. 1999. Stress hormones, Th1/Th2 patterns, pro/anti-inflammatory cytokines and susceptibility to disease. Trends Endocrinol Metab 10: 359-368. Elenkov IJ, Chrousos GP, Wilder RL. 2000a. Neuroendocrine regulation of IL-12 and TNF-alpha/IL-10 balance. Clinical implications. Ann N Y Acad Sci 917: 94-105. Elenkov IJ, Papanicolaou DA, Wilder RL, Chrousos GP. 1996. Modulatory effects of glucocorticoids and catecholamines on human interleukin-12 and interleukin-10 production: Clinical implications. Proc Assoc Am Physicians 108: 374-381. Elenkov IJ, Webster EL, Torpy DJ, Chrousos GP. 1999. Stress, corticotropin-releasing hormone, glucocorticoids, and the immune/inflammatory response: Acute and chronic effects. Ann N Y Acad Sci 876: 1-11. Elenkov IJ, Wilder RL, Bakalov VK, Link AA, Dimitrov MA, et al. 2001. IL-12, TNF-alpha, and hormonal changes during late pregnancy and early postpartum: Implications for autoimmune disease activity during these times. J Clin Endocrinol Metab 86: 4933-4938. Elenkov IJ, Wilder RL, Chrousos GP, Vizi ES. 2000b. The sympathetic nerve – an integrative interface between two supersystems: The brain and the immune system. Pharmacol Rev 52: 595-638. Engstad CS, Lund T, Osterud B. 1999. Epinephrine promotes IL-8 production in human leukocytes via an effect on platelets. Thromb Haemost 81: 139-145. Fearon DT, Locksley RM. 1996. The instructive role of innate immunity in the acquired immune response. Science 272: 50-53. Felten DL, Felten SY, Carlson SL, Olschowka JA, Livnat S. 1985. Noradrenergic and peptidergic innervation of lymphoid tissue. J Immunol 135: 755s-765s. Foreman JC. 1987. Substance P and calcitonin gene-related peptide: Effects on mast cells and in human skin. Int Arch Allergy Appl Immunol 82: 366-371. Fox FE, Kubin M, Cassin M, Niu Z, Hosoi J, et al. 1997. Calcitonin gene-related peptide inhibits proliferation and

18

antigen presentation by human peripheral blood mononuclear cells: Effects on B7, interleukin 10, and interleukin 12. J Invest Dermatol 108: 43-48. Gallucci S, Matzinger P. 2001. Danger signals: SOS to the immune system. Curr Opin Immunol 13: 114-119. Ganea D, Delgado M. 2001. Inhibitory neuropeptide receptors on macrophages. Microbes Infect 3: 141-147. Hasko G, Kuhel DG, Salzman AL, Szabo C. 2000. ATP suppression of interleukin-12 and tumour necrosis factoralpha release from macrophages. Br J Pharmacol 129: 909-914. Hasko G, Szabo C, Nemeth ZH, Kvetan V, Pastores SM, et al. 1996. Adenosine receptor agonists differentially regulate IL-10, TNF-alpha, and nitric oxide production in RAW 264.7 macrophages and in endotoxemic mice. J Immunol 157: 4634-4640. Hasko G, Szabo C, Nemeth ZH, Salzman AL, Vizi ES. 1998. Stimulation of beta-adrenoceptors inhibits endotoxininduced IL-12 production in normal and IL-10 deficient mice. J Neuroimmunol 88: 57-61. Hellstrand K, Hermodsson S. 1989. An immunopharmacological analysis of adrenaline-induced suppression of human natural killer cell cytotoxicity. Int Arch Allergy Appl Immunol 89: 334-341. Ho WZ, Stavropoulos G, Lai JP, Hu BF, Magafa V, et al. 1998. Substance P C-terminal octapeptide analogues augment tumor necrosis factor-alpha release by human blood monocytes and macrophages. J Neuroimmunol 82: 126-132. Hoskin DW, Reynolds T, Blay J. 1994. Adenosine as a possible inhibitor of killer T-cell activation in the microenvironment of solid tumours. Int J Cancer 59: 854-855. Irwin M. 1994. Stress-induced immune suppression: Role of brain corticotropin releasing hormone and autonomic nervous system mechanisms. Adv Neuroimmunol 4: 29-47. Irwin M, Brown M, Patterson T, Hauger R, Mascovich A, et al. 1991. Neuropeptide Y and natural killer cell activity: Findings in depression and Alzheimer caregiver stress. FASEB J 5: 3100-3107. Kaplanski G, Porat R, Aiura K, Erban JK, Gelfand JA, et al. 1993. Activated platelets induce endothelial secretion of interleukin-8 in vitro via an interleukin-1-mediated event. Blood 81: 2492-2495. Karalis K, Sano H, Redwine J, Listwak S, Wilder RL, et al. 1991. Autocrine or paracrine inflammatory actions of corticotropin-releasing hormone in vivo. Science 254: 421-423. Katafuchi T, Take S, Hori T. 1993. Roles of sympathetic nervous system in the suppression of cytotoxicity of splenic natural killer cells in the rat. J Physiol (Lond) 465: 343-357. Kincy-Cain T, Bost KL. 1997. Substance P-induced IL-12 production by murine macrophages. J Immunol 158: 2334-2339.

453

454

18

Sympathetic and peptidergic innervation: Major role at the neural–immune interface

Koff WC, Dunegan MA. 1985. Modulation of macrophagemediated tumoricidal activity by neuropeptides and neurohormones. J Immunol 135: 350-354. Koff WC, Dunegan MA. 1986. Neuroendocrine hormones suppress macrophage-mediated lysis of herpes simplex virus-infected cells. J Immunol 136: 705-709. Kohno M, Kawahito Y, Tsubouchi Y, Hashiramoto A, Yamada R, et al. 2001. Urocortin expression in synovium of patients with rheumatoid arthritis and osteoarthritis: Relation to inflammatory activity. J Clin Endocrinol Metab 86: 4344-4352. la Sala A, Ferrari D, Corinti S, Cavani A, Di Virgilio F, et al. 2001. Extracellular ATP induces a distorted maturation of dendritic cells and inhibits their capacity to initiate Th1 responses. J. Immunol 166: 1611-1617. Le Moine O, Stordeur P, Schandene L, Marchant A, de Groote D, et al. 1996. Adenosine enhances IL-10 secretion by human monocytes. J Immunol 156: 4408-4414. Le Tulzo Y, Shenkar R, Kaneko D, Moine P, Fantuzzi G. et al. 1997. Hemorrhage increases cytokine expression in lung mononuclear cells in mice: Involvement of catecholamines in nuclear factor-kappaB regulation and cytokine expression. J Clin Invest 99: 1516-1524. Lerner BH. 1996. Can stress cause disease? Revisiting the tuberculosis research of Thomas Holmes, 1949–1961. Ann Intern Med 124: 673-680. Levenstein S, Ackerman S, Kiecolt-Glaser JK, Dubois A. 1999. Stress and peptic ulcer disease. JAMA 281: 10-11. Li T, Harada M, Tamada K, Abe K, Nomoto K. 1997. Repeated restraint stress impairs the antitumor T cell response through its suppressive effect on Th1-type CD4+ T cells. Anticancer Res 17: 4259-4268. Li Y, Chi L, Stechschulte DJ, Dileepan KN. 2001. Histamineinduced production of interleukin-6 and interleukin-8 by human coronary artery endothelial cells is enhanced by endotoxin and tumor necrosis factor-alpha. Microvasc Res 61: 253-262. Link AA, Kino T, Worth JA, McGuire JL, Crane ML, et al. 2000. Ligand-activation of the adenosine A2a receptors inhibits IL-12 production by human monocytes. J Immunol 164: 436-442. Liu J, Chen M, Wang X. 2000. Calcitonin gene-related peptide inhibits lipopolysaccharide-induced interleukin-12 release from mouse peritoneal macrophages, mediated by the cAMP pathway. Immunology 101: 61-67. Lotz M, Vaughan JH, Carson DA. 1988. Effect of neuropeptides on production of inflammatory cytokines by human monocytes. Science 241: 1218-1221. Lundberg JM, Rudehill A, Sollevi A, Fried G, Wallin G. 1989. Co-release of neuropeptide Y and noradrenaline from pig spleen in vivo: Importance of subcellular storage, nerve

impulse frequency and pattern, feedback regulation and resupply by axonal transport. Neuroscience 28: 475-486. Maestroni GJ. 1998a. Is hematopoiesis under the influence of neural and neuroendocrine mechanisms?. Histol Histopathol 13: 271-274. Maestroni GJ. 1998b. Catecholaminergic regulation of hematopoiesis in mice [letter; comment]. Blood 92: 2971-2973. Maisel AS, Michel MC. 1990. Beta-adrenoceptor control of immune function in congestive heart failure. Br J Clin Pharmacol 30 (Suppl 1): 49S-53S. Marshall GD, Jr, Agarwal SK. 2000. Stress, immune regulation, and immunity: Applications for asthma. Allergy Asthma Proc 21: 241-246. Matzinger P. 2002. The danger model: A renewed sense of self. Science 296: 301-305. Miles BA, Lafuse WP, Zwilling BS. 1996. Binding of alphaadrenergic receptors stimulates the anti-mycobacterial activity of murine peritoneal macrophages. J Neuroimmunol 71: 19-24. Miller LE, Justen HP, Scholmerich J, Straub RH. 2000. The loss of sympathetic nerve fibers in the synovial tissue of patients with rheumatoid arthritis is accompanied by increased norepinephrine release from synovial macrophages. FASEB J 14: 2097-2107. Mohamed-Ali V, Flower L, Sethi J, Hotamisligil G, Gray R, et al. 2001. Beta-adrenergic regulation of IL-6 release from adipose tissue: In vivo and in vitro studies. J Clin Endocrinol Metab 86: 5864-5869. Mosmann TR, Sad S. 1996. The expanding universe of T-cell subsets: Th1, Th2 and more. Immunol Today 17: 138-146. Munford RS, Tracey KJ. 2002. Is severe sepsis a neuroendocrine disease? Mol Med 8: 437-442. Murray DR, Prabhu SD, Chandrasekar B. 2000. Chronic betaadrenergic stimulation induces myocardial proinflammatory cytokine expression. Circulation 101: 2338-2341. Nair MP, Schwartz SA, Wu K, Kronfol Z. 1993. Effect of neuropeptide Y on natural killer activity of normal human lymphocytes. Brain Behav Immun 7: 70-78. O’Sullivan ST, Lederer JA, Horgan AF, Chin DH, Mannick JA, et al. 1995. Major injury leads to predominance of the T helper-2 lymphocyte phenotype and diminished interleukin12 production associated with decreased resistance to infection [see comments]. Ann Surg 222: 482-490. Padgett DA, Sheridan JF, Dorne J, Berntson GG, Candelora J, et al. 1998. Social stress and the reactivation of latent herpes simplex virus type 1. Proc Natl Acad Sci U S A 95: 72317235. Panina-Bordignon P, Mazzeo D, Lucia PD, D’Ambrosio D, Lang R, et al. 1997. Beta2-agonists prevent Th1 development by selective inhibition of interleukin 12. J Clin Invest 100: 1513-1519.

Sympathetic and peptidergic innervation: Major role at the neural–immune interface Perregaux DG, McNiff P, Laliberte R, Conklyn M, Gabel CA. 2000. ATP acts as an agonist to promote stimulus-induced secretion of IL-1 beta and IL-18 in human blood. J. Immunol 165: 4615-4623. Prabhakar U, Brooks DP, Lipshlitz D, Esser KM. 1995. Inhibition of LPS-induced TNF alpha production in human monocytes by adenosine (A2) receptor selective agonists. Int J Immunopharmacol 17: 221-224. Rook GA, Lightman SL, Heijnen CJ. 2002. Can nerve damage disrupt neuroendocrine immune homeostasis? Leprosy as a case in point. Trends Immunol 23: 18-22. Sainz B, Loutsch JM, Marquart ME, Hill JM. 2001. Stressassociated immunomodulation and herpes simplex virus infections. Med Hypotheses 56: 348-356. Sanders VM, Baker RA, Ramer-Quinn DS, Kasprowicz DJ, Fuchs BA, et al. 1997. Differential expression of the beta2adrenergic receptor by Th1 and Th2 clones: Implications for cytokine production and B cell help. J Immunol 158: 4200-4210. Sanders VM, Munson AE. 1985. Norepinephrine and the antibody response. Pharmacol Rev 37: 229-248. Shakhar G, Ben-Eliyahu S. 1998. In vivo beta-adrenergic stimulation suppresses natural killer activity and compromises resistance to tumor metastasis in rats. J Immunol 160: 3251-3258. Spengler RN, Allen RM, Remick DG, Strieter RM, Kunkel SL. 1990. Stimulation of a-adrenergic receptor augments the production of macrophage-derived tumor necrosis factor. J Immunol 145: 1430-1434. Sperlagh B, Doda M, Baranyi M, Hasko G. 2000. Ischemic-like condition releases norepinephrine and purines from different sources in superfused rat spleen strips. J Neuroimmunol 111: 45-54. Straub RH, Cutolo M. 2001. Involvement of the hypothalamic– pituitary–adrenal/gonadal axis and the peripheral nervous system in rheumatoid arthritis: Viewpoint based on a systemic pathogenetic role. Arthritis Rheum 44: 493-507. Straub RH, Herrmann M, Frauenholz T, Berkmiller G, Lang B, et al. 1996. Neuroimmune control of interleukin-6 secretion in the murine spleen. Differential beta-adrenergic effects of electrically released endogenous norepinephrine under various endotoxin conditions. J Neuroimmunol 71: 37-43. Straub RH, Mayer M, Kreutz M, Leeb S, Scholmerich J, et al. 2000. Neurotransmitters of the sympathetic nerve terminal are powerful chemoattractants for monocytes. J Leukoc Biol 67: 553-558. Theoharides TC, Singh LK, Boucher W, Pang X, Letourneau R, et al. 1998. Corticotropin-releasing hormone induces skin mast cell degranulation and increased vascular permeability, a possible explanation for its proinflammatory effects. Endocrinology 139: 403-413. 93.

18

Theoharides TC, Spanos C, Pang X, Alferes L, Ligris K, et al. 1995. Stress-induced intracranial mast cell degranulation: A corticotropin-releasing hormone-mediated effect, Endocrinology 136: 5745-5750. Trinchieri G. 1995. Interleukin-12: A proinflammatory cytokine with immunoregulatory functions that bridge innate resistance and antigen-specific adaptive immunity. Annu Rev Immunol 13: 251-276. Udelsman R, Gallucci WT, Bacher J, Loriaux DL, Chrousos GP. 1986. Hemodynamic effects of corticotropin releasing hormone in the anesthetized cynomolgus monkey. Peptides 7: 465-471. van der Poll T, Coyle SM, Barbosa K, Braxton CC, Lowry SF. 1996. Epinephrine inhibits tumor necrosis factor-alpha and potentiates interleukin 10 production during human endotoxemia. J Clin Invest 97: 713-719. Vicennati V, Vottero A, Friedman C, Papanicolaou DA. 2002. Hormonal regulation of interleukin-6 production in human adipocytes. Int J Obes Relat Metab Disord 26: 905-911. Vizi ES, Orso E, Osipenko ON, Hasko G, Elenkov IJ. 1995. Neurochemical, electrophysiological and immunocytochemical evidence for a noradrenergic link between the sympathetic nervous system and thymocytes. Neuroscience 68: 1263-1276. Vredevoe DL, Moser DK, Gan XH, Bonavida B. 1995. Natural killer cell anergy to cytokine stimulants in a subgroup of patients with heart failure: Relationship to norepinephrine. Neuroimmunomodulation 2: 16-24. Weihe E, Nohr D, Michel S, Muller S, Zentel HJ, et al. 1991. Molecular anatomy of the neuro-immune connection. Int J Neurosci 59: 1-23. Westfall DP, Todorov LD, Mihaylova-Todorova ST. 2002. ATP as a cotransmitter in sympathetic nerves and its inactivation by releasable enzymes. J Pharmacol Exp Ther 303: 439-444. Whalen MM, Bankhurst AD. 1990. Effects of beta-adrenergic receptor activation, cholera toxin and forskolin on human natural killer cell function. Biochem J 272: 327-331. Wilder RL. 1995. Neuroendocrine–immune system interactions and autoimmunity. Annu Rev Immunol 13: 307-338. Wilkin F, Stordeur P, Goldman M, Boeynaems JM, Robaye B. 2002. Extracellular adenine nucleotides modulate cytokine production by human monocyte-derived dendritic cells: Dual effect on IL-12 and stimulation of IL-10. Eur J Immunol 32: 2409-2417. Woiciechowsky C, Asadullah K, Nestler D, Eberhardt B, Platzer C, et al. 1998. Sympathetic activation triggers systemic interleukin-10 release in immunodepression induced by brain injury [in process citation]. Nat Med 4: 808-813.

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Index

AADC. See Aromatic L-amino acid decarboxylase Acetylcholine, 102–105, 107, 109 – breakdown, 115, 116 – depot, 115 – regulation by – 5-HT1A receptors, 177 – 5-HT6 receptors, 192 – regulation of – release by 5-HT3 receptors, 187, 189 – release by 5-HT4 receptors, 189, 190 – release, 114–116, 120 – synthesis, 114, 115 – vesicle pool, 114 Acetylcholine-esterase (AChE), 115, 116 Acetylcholine (ACh) neurons – ambient level of Ach, 10 – asynaptic features, 9, 13, 14 – axon varicosities (terminals), 8, 9, 13 – basalocortical ACh system, 8–10 – cell groups, 10, 11 – developmental aspects, 13 – diffuse transmission, 9, 10 – innervation of – hippocampus, 8, 10, 13 – neocortex, 8–10, 13 – neostriatum, 8, 10, 13 – spinal cord, 8, 11 – interneurons, 8–10, 13 – major projections, 8 – septohippocampal ACh system, 10, 13 – synaptic features, 9–11, 13 Acetyl coenzyme A, 114, 115 Addiction – links to 5-HT1B receptors, 173 – role of 5-HT3 receptors, 189 Adenosine, 229–235, 237–239, 241, 330, 331, 333 #

– receptors, 258–260, 265 – release, 264 – transporters, 257 Adenosine 50 -diphosphate, ADP, 229–231, 234, 235, 237, 238, 245 Adenosine 50 -monophosphate, AMP, 229–231, 234, 235 Adenosine receptors – regulation of AMPA receptors by, 279, 300 Adenosine 50 -triphosphate, ATP, 228–239, 241–246 Adenylate cyclaseb (AC) – 5-ht5a receptor coupling, 190 – inhibition by – 5-HT1A receptors, 175 – 5-HT1B receptors, 179 – 5-HT1D receptors, 181 – 5-HT1E receptors, 182 – regulation by – adenosine receptors, 300 – adrenergic receptors, 300, 301 – metabotropic glutamate receptors, 287 – opioid receptors, 301 – serotonin receptors, 298, 300 – stimulation by – 5-HT4 receptors, 189 – 5-HT6 receptors, 192 – 5-HT7 receptors, 194 Adenylate kinase, 230, 234 Adrenaline neurons – anatomical distribution, 6 – cell groups, 6 – in spinal cord, 6 Adrenergic receptors, regulation of AMPA receptors by, 300, 301 AEA membrane transporter’ (AMT), 344, 353, 358–360, 363 2-AG, 344, 353–368 Aggregated Ab42, direct vs. indirect neurotoxicity, 392

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Aggression – links to 5-HT1A receptors, 177 – links to 5-HT1B receptors, 179 Alcohol – non-preferring, 369 – preferring, 369 – related phenomena, 368 Alcohol tolerance, vulnerability, reinforcement, and consumption, 368 Alcohol withdrawal symptoms, 369, 370 Alkaline phosphatase, 230, 234 Allergy, 450, 451 Alternative splicing – 5-HT4 receptors, 174 – 5-HT7 receptors, 194, 197 Alzheimer’s disease, 114, 115, 121, 122, 256, 270, 271, 344, 368, 392, 395 – role of 5-HT1A receptors, 178 – role of 5-HT4 receptors, 190 AM‐281, 365 a-amino‐3-hydroxy‐5-methyl‐4isoxazole propionate (AMPA) – receptor, 416, 417, 421, 427, 430 g-aminobutyric acid, 46 – attenuation by 5-HT1A receptors, 175 AMPA currents AMPA receptor expression (AMPAR expression) – effect of – antipsychotic drugs on, 297 – drugs of abuse on, 293, 297 – regulation by – dopamine, 279, 288, 290–308 – receptor tyrosine kinases, 277, 278, 284, 303, 308 – serotonin, 279, 298, 300 AMPA receptor interacting proteins – general AMPA receptor interactors: transmembrane and extracellular proteins, 286

458

Index – proteins that interact with long forms of AMPA receptor subunits, 281, 284–286 – proteins that interact with short forms of AMPA receptor subunits, 285, 286 AMPA receptor phosphorylation – PKA mediated, 283 – PKC mediated, 283 – regulation by – adrenergic receptors, 283 – insulin, 284 – metabotropic glutamate receptors, 284 – serotonin, 298 – role in – LTD, 283, 284 – LTP, 283 – receptor trafficking, 284 AMPA receptor structure – glutamate-binding domain, 281 – regulation by – alternative splicing, 278, 281 – RNA editing, 278, 281 – topology of AMPA receptors, 281, 282 AMPA receptor trafficking, 284 AMT. See AEA membrane transporter Amygdala – 5-HT1A receptor, 175 – 5-HT1B receptor location, 179 – 5-HT1E receptor location, 182 – 5-HT2C receptor location, 186 – 5-HT3 receptor location, 187 – 5-HT5A receptor location, 190 – 5-HT7 receptor location, 194 – SERT location, 196–198 Amyloid lateral sclerosis – COX‐2, 396 – familial form mutation, 394, 396 Analgesia – 5-HT3 receptor ligand potential, 173, 174, 187 Ananda, ‘‘internal bliss’’, 356 Anandamide, 344–346, 353, 356–359, 361, 363, 364 Anandamide hydrolase, 357 Animal models of neurodegenerative diseases, 386, 392, 394, 396 Anorexia nervosa, role of 5-HT2A receptors, 185 Anticholinesterases, 115 Anxiety – 5-HT1B receptor role, 179 – 5-HT2B receptor role, 185 – role of – 5-HT2C receptors, 186

– 5-HT3 receptors, 188 – 5-HT4 receptors, 190 – 5-HT6 receptors, 193 – SERT, 197, 198 2-Arachidonylglycerol, 345, 356–359 2-Arachidonylglycerol ether, 345, 356 Arcuate nucleus, TH, DAT and VMAT2 expression, 45 Area postrema – location of 5-HT3 receptors, 187 Aromatic L-amino acid decarboxylase, 23 Aromatic L-amino acid decarboxylaseimmunoreactive neurons, distribution of, 60 Astrocytes, 263, 264, 267 Astrocytic Ca2+ waves, 241 Atopy, 451 ATP, 256–258, 262–267 Autoimmunity, 446 Autoradiography, 76, 77, 82, 83 – distribution of – 5-HT1A receptors, 175 – 5-HT1B receptors, 178 – 5-HT1D receptors, 181 – 5-HT1F receptors, 182 – 5-HT2A receptors, 183 – 5-HT2C receptors, 186 – 5-HT4 receptors, 189 – 5-HT7 receptors, 194 – SERT, 196 Basal ganglia – 5-HT1B receptor location, 179 – 5-HT1D receptor location, 181 – 5-HT4 receptor location, 189 – SERT location, 196 BDNF. See Brain-derived neurotrophic factor Benzodiazepines, 219 Biacylglycerol, 344, 358, 359 Bienzymatic neurons, 27 Bienzymatic TH-and AADC-expressing neurons, in Parkinson’s disease, 59 Biosynthesis and metabolism, 357, 359 Bipolar disorder, 130, 135, 136, 139, 140 Brain cannabinoid receptor, 346, 352 Brain-derived neurotrophic factor – activation by 5-HT2A receptors, 183, 184 Brain noradrenergic system – nuclei and pathways of, 131 – ontogeny of, 130, 131 Brainstem, medulla, limbic forebrain, striatum, and C57BL/6 mice, 350, 369

Bulimia nervosa, role of 5-HT2A receptors, 185 g‐Butyrolactone (GBL) model, 162 Ca2+-and TTX-sensitive, 366 Caffeine, 258–261, 269–271 Calcium – dependency, 331 – homeostasis, 326, 334 Calcium/calmodulin-dependent protein kinase II (CaMKII), 427–430, 432 Calcium-dependent transacylase, 344, 357, 363 Cannabinoid signaling, 355, 362, 365, 368, 369 Cannabinoids have specific receptors, 346 Carbon monoxide – effector mechanisms, 412 – synthesis of CO, 412 Catecholamines, 23 – epinephrine, 444–449, 451 – norepinephrine, 444, 449, 451 Catechol-O-methyl-transferase (COMT), 155 Caudate putamen – 5-HT1D receptor transcript location, 181 – 5-ht1E receptor location, 182 – 5-HT1F receptor location, 182 – 5-HT2A receptor location, 183 – 5-HT2C receptor location, 186 – 5-HT3 receptor location, 187 – 5-HT4 receptor location, 189 – 5-HT6 receptor location, 192 CB1 and CB2 receptor knockout mice, 349 CB1 receptor, 344, 346–362, 364, 370, 371 – agonist, 371 – antagonist, 371 – knockout, 350 CB2, 344–358 CB3, anandamide receptors, 346, 364, 366 Cerebellum, 352, 357, 359–361, 364, 365, 367 – 5-HT1B receptor location, 179 – 5-HT2C receptor location, 186 – 5-HT5A receptor location, 190 – 5-HT6 receptor location, 192 Chemosensory transduction, 243 Chloride conductance, 332, 334 Choline, 114, 115, 117, 122 – transport, 114 – uptake, 114, 115

Index Choline-acetyltranspherase (ChAT), 114, 115 Cholinergic – interneurons, 116 – projection neurons, 116 – transmission, 114–116, 118, 119 Cholinergic neurones – 5-HT1A receptors, 175 Cholinergic receptors – regulation of AMPA receptors by, 300 – signaling mechanisms, 296, 300 Choroid plexus, location of 5-HT2C receptors, 186 Chronic alcohol, 359, 368–370 Circadian rhythm – links to – 5-HT1A receptors, 178 – 5-HT1B receptors, 181 – role of – 5-HT5A receptors, 191 – 5-HT7 receptors, 194, 195 Claustrum – 5-HT1E receptor location, 182 – 5-HT1F receptor location, 182 Cortex – 5-HT1D receptor location, 181 – 5-HT1E receptor location, 182 – 5-HT1F receptor location, 182 – 5-HT2A receptor location, 183 – 5-HT2C receptor location, 186 – 5-HT4 receptor location, 189 – 5-HT5A receptor location, 190 – 5-HT6 receptor location, 192 – 5-HT7 receptor location, 194 – SERT location, 196, 197 COXs – COX‐1, 395 – COX‐2, 392, 394–396 CP‐55, 940, 344 cPLA2, 344, 354 CSF, increased PGE2, 394, 396 Cytokines, 245 – anti-inflammatory cytokines, 446, 447 – proinflammatory cytokines, 444, 446, 447, 450 Cytolysis, 241–243, 245 DA, cooperative synthesis of, 49, 50 D2 DA receptors, 366 DA-ergic phenotype expression – striatal neurons in, 53, 54 – tuberoinfundibular neurons in, 52, 53 DA synthesis in monoenzymatic AADC-neurons, 47 DAT. See Dopamine transporter

DA transporter expression, 35 DBA/2 mice, 369 Depression, 130, 134, 135 – links to – 5-HT1A receptors, 175, 177 – 5-HT1B receptors, 179 – 5-HT2A receptors, 183 – role of – 5-HT2C receptors, 185 – 5-HT6 receptors, 193 – 5-HT7 receptors, 196 – SERT, 197, 198 Development, 421–423 DHPG, 344, 365 Diacylglycerol lipase, 359 Diagonal band of Broca – 5-HT1A receptors, 175 Diencephalon, TH-immunoreactive neurons in, 46 3,4-Dihydroxyphenylacetic acid (DOPAC), 155, 157 Displacement, drug abuse, 89 Dopamine – as classic neurotransmitters, 152 – degradative pathway of, 155 – regulation by – 5-HT1A receptors, 175 – 5-HT2A receptors, 183 – 5-HT2C receptors, 186 – 5-HT6 receptors, 192 – release due to 5-HT3 receptors, 187 – release facilitation by 5-HT4 receptors, 189 Dopamine (DA) neurons – ambient level, 4 – asynaptic features, 4 – axon morphology, 3, 4 – cell groups, 2–4 – content of single varicosity, 3 – developmental aspects, 11, 12 – diencephalospinal DA system, 4 – diffuse transmission by, 4 – innervation of – cerebral cortex, 3 – neostriatum, 4 – spinal cord, 3, 4 – major projections, 2–4 – mesocortical DA system, 3, 5 – mesostriatal DA system, 2–4 – morphometric data, 3 – synaptic features, 3, 4, 12 Dopamine neurotransmission, 333 Dopamine plasma membrane transporter (DAT) – drugs action on, 159 – operation of, 159 – structure of, 158

Dopamine receptors – postsynaptic, 161 – presynaptic, 161–162 – regulation of – AMPA receptor activity by, 296, 297 – AMPA receptor phosphorylation by, 291, 297 – AMPA receptor surface expression by, 279 – role in synaptic plasticity, 293, 308 – role of DARPP–32, 292, 293, 299, 300 – structure of, 160, 161 – supersensitivity of, 162, 163 – types of, 160 Dopamine release – action potential propagation induced, 163, 164 – by ion channel-coupled receptors, 165 – by reverse-mode operation, 164 – regulation of, 165 Dopamine, synthetic pathways of, 24 Dopamine transporter, 45 Dopaminergic tuberoinfundibular and nigrostriatal systems, in rat, 53 Dorsal motor nucleus – location of 5-HT3 receptors, 187 Dorsal periventricular pathway, 172 Dorsal raphe nucleus – 5-HT1B receptor transcript location, 179 – 5-HT1D receptor transcript location, 181 – 5-HT5a receptor location, 190 – 5-HT7 receptor location, 194 Down-regulation, 368, 369 Drink less alcohol, 369, 370 DSE, 365, 366 DSI, 344, 365–367 EctoATPase, 234 Ecto-nucleotidases, 263, 264, 267 Effect of H2O2 on potassium channels, 412 Electrophysiology, of 5-HT neurons, 172 Emesis, role of 5-HT3 receptor ligands, 187, 188 Endocannabinoids, 345–367 Energy charge, 257 Entorhinal cortex – 5-HT1A receptors, 175 – 5-ht5b receptor transcript location, 192

459

460

Index EP receptors – adaptive immune response, 388, 392 – excitotoxicity, 392, 393, 395 – expression, 387, 388, 390, 392, 393 – innate immune response, 392, 393, 395 – multiple subtypes, 386–389 – periphery, 388 EP1 – excitotoxicity, 392, 395 EP2 – EP2
/
, 393 – innate immunity, 388 – paracrine damage, 393, 395 – periphery, 388 EP4 – excitotoxicity, 392 Epilepsy – role of – 5-HT6 receptors, 194 – 5-HT7 receptors, 196 – SERT, 198 EPSCs, 366, 367 Excitotoxicity, 266–268 – COX‐2 neuronal expression, 391 – N-Methyl-D-Aspartic Acid (NMDA) receptor, 391 Exocytosis, 331 Extracellular space, 103, 106–108 FAAH, 344, 352, 353, 357–363, 368, 371 FAAH-immunoreactivity, 362 FAAH-KO mice, 361, 362 Facilitated diffusion, 363 Fast excitatory neurotransmission, 232, 238, 239, 241, 243 Fatty acid amide (FAA) metabolism, 359 Feeding, role of – 5-HT2B receptors, 185 – 5-HT2C receptors, 186 Four clusters of Ca2+-calmodulindependent protein kinase sites, 349 Four clusters of cAMP-dependent protein kinase sites, 349 Frontal cortex, modulation of 5-HT release by 5-HT1B receptors, 179 GABA, 344, 352, 357, 362, 364–366 See also g-aminobutyric acid – degradation, 216, 217 – metabolism, 216, 217 – receptors, 219–220, 329, 330, 332 – regulation by – 5-HT6 receptors, 192 – 5-HT7 receptors, 194, 195 – release, 218–219

– release due to 5-HT2A receptors, 183, 184 – synthesis, 214–216 – transport, 218, 219, 221–222 GABA, extrasynaptic, 106 GABAA receptor agonists – GABA, 220 – isoguvacine, 220 – isonipecotic acid, 220 – muscimol, 220 – THIP, 219 GABAA receptor antagonists – bicuculline, 219, 220 – picrotoxinin, 220 GABA-shunt, 216 GABA-T, 216 GABA transport inhibitors – EF 1502, 222 – exo-THPO, 222 – guvacine, 221 – nipecotic acid, 221 – THPO, 221 – tiagabine, 222 GABAergic interneurones – 5-HT1A receptors, 175 – expression of 5-HT2A receptors, 183 GAD. See Glutamic acid decarboxylase Gene expression, 354, 368–370 Genes are located in the proximal arm of chromosome 4, 352 Glial cells, 327, 328 Gliotransmission, 241 Globus pallidus – location of 5-HT4 receptors, 189 Glutamate, 105, 107, 344, 352, 357, 362, 364–367 – neurotransmitter, 426, 430 – regulation by 5-HT6 receptors, 192 – release due to 5-HT2A receptors, 183 – spillover, 426 Glutamate receptor – agonist, 416–418, 434 – antagonist, 417, 418, 421 – distribution, 423, 424, 427, 434 – editing, 418, 420–423 – ER retention, 433 – genes, 416, 420 – glutamate receptor interacting protein 1 (GRIP1), 416, 427 – hippocampal neuron, 421, 423, 433 – ionotropic, 416, 417, 421 – isoforms, 416–418, 426

– kainate receptor, 417, 422, 424, 429, 432, 433 – metabotropic, 416, 420, 421, 423 – pharmacological, 416, 417, 420, 426 – phosphorylation, 416, 417, 427–430, 432–434 – protein-protein interaction, 416, 417, 427, 434 – regulation, 421–423, 427, 429 – splice variants, 417, 418, 420, 434 – structure, 417, 421 – subunits, 416–418, 420–424, 427, 428, 430, 432, 433 – targeting, 421–423, 430, 431, 433, 434 Glutamic acid decarboxylase, 214–216 Glycine receptors, 328, 329, 332, 333 G protein-coupled receptors, 132, 133 G-protein modulated potassium current – modulation by 5-HT1A receptors, 175 G-proteins, 287, 300–302 Granule cells – 5-HT1A receptors, 175, 177 GTP. See Guanosine triphosphate Guanosine triphosphate (GTPS), 44, 344 Hemicholinium, 114 Heterosynaptic depression, 266, 267 Highest densities present in the, substantia nigra pars reticulata, 352 Hippocampus, 352, 354, 357, 359–370 – and cerebellum, 352, 357–361, 364, 365 – 5-HT1A receptor localization, 175, 177, 178 – 5-HT1B receptor location, 179 – 5-HT1D receptor location, 181 – 5-HT1E receptor location, 182 – 5-HT2A receptor location, 183, 185 – 5-HT2C receptor location, 186, 187 – 5-HT3 receptor location, 187, 189 – 5-HT4 receptor location, 189 – 5-HT5A receptor location, 190 – 5-HT5B receptor transcript location, 190 – 5-HT6 receptor location, 192, 193 – 5-HT6 receptor transcript location, 192 – 5-HT7 receptor location, 194, 196 – modulation of – acetylcholine release by 5-HT1B receptors, 179 – 5-HT release by 5-HT1B receptors, 179

Index – release of NA by 5-HT1A receptors, 175 – SERT location, 196–198 Histamine neurons – anatomical distribution, 7, 8 – asynaptic features, 8 Homeostasis, 257 Homodimers/Heterodimers – 5-HT4 receptors and b2 adrenoceptors, 174, 189 Homovanillic acid (HVA), 155 5-HT – regulation of release by 5-HT7 receptors, 194 – release due to 5-HT3 receptors, 187–189 – release due to 5-HT4 receptors, 189, 190 5-HT moduline – modulation of 5-HT1B receptor binding, 181 5-HT7 receptor location, 194–196 Huntington’s disease, 344, 367, 368 Hydrogen peroxide – synthesis of H2O2, 412 Hyperpolarization, 328, 329, 331, 334 Hyperprolactinemia, DA-ergic phenotype in, 52, 53 Hypolocomotive, 349 Hypotensive, 349 Hypothalamic magnocellular vasopressinergic neurons, 44 Hypothalamic-pituitary-adrenal axis, 131 Hypothalamus – 5-HT1A receptors, 175 – 5-HT1B receptor location, 178–181 – 5-HT7 receptor location, 196 – location of 5-ht5a receptor, 190–192 – role of 5-HT2A receptor in secretion of hormones, 185 – SERT location, 196, 197 Hypothermia, role of 5-HT7 receptors, 194–196 Hypothermic effects of cannabinoids, 349 Immunohistochemistry – 5-HT1A receptor, 175 – distribution of 5-HT1B receptors, 178 – location of – 5-HT2B receptors, 185 – 5-HT2C receptors, 186 – 5-ht5a receptors, 190 – 5-HT6 receptors, 192–194 – SERT, 196

Impulse-mediating dopamine autoreceptor, 162 In situ hybridisation – 5-HT1A receptors, 175 – distribution of – 5-HT1B receptors, 178, 179 – 5-HT1D receptors, 181 – 5-HT1E receptors, 182 – 5-HT2C receptors, 186 – 5-HT5A receptors, 190 – 5-HT5B receptors, 192 – 5-HT6 receptors, 192 – 5-HT7 receptors, 194 – SERT, 196–198 Inhibit acute alcohol-induced dopamine release, 350 Inhibit alcohol consumption, 369, 370 Innate immune response in brain – LPS, 393 – major effector in several chronic neurodegenerative diseases, 392 – paracrine oxidative damage to neurons, 392, 393 – phagocytosis of deleterious protein aggregates, 392, 393 Innervation – parasympathetic (cholinergic), 444 – peptidergic/sensory, 444, 445, 449 – sympathetic, 444, 445, 447, 450, 451 Inosine, 229, 230, 234, 235 Interleukins – IL–1, 444, 451 – IL–2, 446 – IL–4, 447, 450 – IL–6, 444, 451 – IL–10, 444, 447, 448, 452 – IL–12, 444, 447, 448, 452 – IL–18, 444, 447 Internal and external segments of the globus pallidus, 352 Ionotropic receptors, 219 IPSCs, 365, 366 Ischemia, 231, 232, 234, 244, 245 – COX‐2, 394, 395 – EP1, 395 JNK, 344, 354 Knockout mouse – 5-HT1A receptors, 177, 178 – 5-HT1B receptors, 179–181 – 5-HT2A receptors, 183, 184 – 5-HT2C receptors, 186 – 5-ht5a receptors, 191 – 5-HT7 receptors, 195, 196 – SERT, 197 L–3,4-dihydroxyphenylalanine (L-DOPA), 23

L-amino

acid decarboxylase (AADC), 23 – enzymatic activity of, 25 – immunoreactive neurons, in rats, 35 – monoenzymatic, 32 L-Aromatic amino acid carboxylase (AADC), 154 L-DOPA – AADC role in, 47 – administration of, 60, 61 – concentration in – AN of fetal rats, 49 – mediobasal hypothalamus, 44 – inhibition of, 50 – L-tyrosine stimulates, 51 – role in disease treatment, 55 – role of monoenzymatic TH-neurons in, 46–47 – synthesis in monoenzymatic TH-neurons, 46, 51 L-tyrosine conversion, 23 Lack highly conserved proline residues, 348 Larger long-term potentiation (LTP), 344, 350, 367 Levuglandins, 387 Lipophilicity, 84, 85 Localization and distribution, 357, 362 Locus coeruleus, 131, 134, 135 – 5-HT1D receptor transcript location, 181 – 5-ht5a receptor location, 190 – SERT location, 196 Long term depression (LTD), 344, 367, 427 Long term potentiation (LTP), 264, 265, 422 LOXs, 386 LPS – co-receptors CD14 and TLR4, 393 – effects in Brain, 393 – EP2, 393, 395 – NSAIDs, 393 Macrophages, 388 MA-ergic neurons, 25 MA-ergic phenotype expression, 25 – brain neurons expressions, in adult mammals – bienzymatic TH-and AADC-expressing neurons, 26–32 – individual enzymes expression, MA synthesis, 26–32 – non-MA-ergic neurons expression, 33–35

461

462

Index – brain neurons expressions, in mammals ontogenesis – individual enzymes expression, MA synthesis, 35–42 – non-MA-ergic neurons expression, 42, 43 – functional properties, 44 – ensembles of monoenzymatic neurons, 48–51 – monoenzymatic neurons expressing AADC, 47, 48 – monoenzymatic neurons expressing TH, 44–47 – non-MA-ergic neurons, 51, 52 – neurons discovery, 26 – regulation of – by diffusive factors, 64, 65 – by neural afferents, 62–64 – hormonal regulation of, 65, 66 Mast cells, 445, 446, 449 Mechanism of inhibition of neurotransmitter release, 366 Medial forebrain bundle, 172 Medial habenula – 5-HT2C receptor location, 186 Median raphe nucleus – 5-HT1B receptor transcript location, 179 – 5-ht5a receptor location, 190 Mediate the analgesic, 349 Memory & learning – links to 5-HT1A receptors, 178 – role of – 5-HT2A receptors, 185 – 5-HT2C receptors, 187 – 5-HT3 receptors, 189 – 5-HT4 receptors, 190 – 5-HT6 receptors, 193 – 5-HT7 receptors, 196 mEPSCs, 366 Metabolism – catechol-O-methyltransferase, 134, 139, 140 – dopamine-b-hydroxylase, 131, 137, 138 – mono-amine-oxydase, 134, 138, 139 – tyrosine hydroxylase, 131, 136, 137 Metabotropic glutamate receptors – internalization of AMPA receptors induced by, 288–290 – regulation of AMPA receptors by, 288–290

– role in synaptic plasticity, 281, 283, 301, 308 – signaling mechanisms, 298, 303 Metabotropic glutamate receptors, 330 Metabotropic receptors, 220 1-methyl–4-phenyl–1,2,3,6tetrahydropyridine (MPTP), 55, 57, 58, 167 a
methyl-p-tyrosine, dopamine turnover rate on, 154 Microglia, 263, 269 Migraine – role of – 5-HT1F receptors, 182 – 5-HT2B receptors, 185 – treatment with – 5-HT1B receptor ligands, 181 – 5-HT1D receptor ligands, 182 Missense mutation, 363 Monoacylglycerol, 358, 360, 363 Monoamine oxidase (MAO), 155 Monoamines (MAs), 23 – in secretory granules, 25 Monoenzymatic AADC-neurons, 47, 48 Monoenzymatic TH neurons, 44–47 Mouse Models of AD – Amyloid Precursor Protein (APP), 395 – EP2, 388, 392, 393 – EP2
/
, 392, 393 – NSAIDs, 395 MPTP. See 1-methyl‐4-phenyl‐1,2,3,6tetrahydropyridine Muscarinic acetylcholine receptors (mAChRs) – functions, 122 – mechanisms, 121 – pharmacology, 122 – subunits and subtypes, 120, 121 Muscarinic acetylcholine receptors, 367 NAPE-specific PLD, 357 N-arachidonyl-dopamine, 344 N-arachidonylethanolamine (AEA) and, 345, 354, 357 N-ArPE, 344, 357 Natriuretic peptide receptors – regulation of AMPA receptors by, 302 – signaling mechanisms, 302, 303 Negatively coupled to adenylate cyclase, 355 Nerve terminals, 261–264, 270 N-ethylmaleimide-sensitive factor (NSF), 416, 432 Neural inhibition, 326, 329, 332

Neuroimaging – ex vivo, 76 – functional, 76 – in vitro, 76 – post mortem, 76 Neuroimmunomodulation, 228, 245, 246 Neuroinflammation, 269 Neuromodulation, 260, 261, 263, 266, 271 – definition, 2 – diverse modes of neuronal communication, 14 – extrinsic, 14 – intrinsic, 14 – neuromodulators in brain, 2 Neuromodulator(s), 326, 331, 334, 357 Neuron-glia communication, 241 Neurons, 327–332, 334 Neuropeptides, 444, 445, 449 – CGRP, 444, 445, 448, 449 – NPY, 444–448, 450 – substance P, 444, 445, 448, 449, 451 – VIP, 444, 445 Neuroprotection, 245 Neuropsychiatric disorder – ADHD, 89 – Huntington’s disease, 89, 95 – Parkinson’s disease, 89, 93 – Schizophrenia, 76, 89, 93 Neuroreceptor – dopamine, 76, 77, 87, 89, 91, 93 – endocannabinoid, 95 – GABA-benzodiazepine, 91, 93 – glutamate, 90, 94 – muscarinic, 90, 93 – nicotinic, 90, 93 – norepinephrine, 90, 93 – opioid, 95 – peripheral benzodiazepine, 94 – serotonin, 87, 90, 91 Neurotransmitter(s)/ neurotransmission, 326, 328–331, 333, 334, 444, 449, 450 – acetylcholine, 93 – ATP, 444, 445, 447, 448, 450 – dopamine, 76, 77, 87, 89, 91, 93 – dopaminergic, 89 – epinephrine, 444–449, 451 – glutamate, 90 – hydroxytryptamine, 91 – monoaminergic, 91, 93 – norepinephrine, 90, 93, 444, 449, 451 – serotonin, 87, 90, 91

Index Nicotinic acetylcholine receptors (nAChRs) – Ca2+/Na+ permeability, 117 – desensitization, 117, 118 – in plasticity, 119, 120 – in reward, 119 – pre-/postsynaptic, 118, 119 – selective ligands, 117 – subunits and subtypes, 117 Nitric oxide – effector mechanisms – binding to haem groups, 407, 408 – S-nitrosylation, 407 – role of NO in the CNS, 408 – NO and LTP, 408, 409 – NO and neurotoxicity, 410, 411 – NO and nonsynaptic interactions, 409, 410 – role of NO in the peripheral nervous system, 411 – synthesis of NO – inhibitors, 407 – nitric oxide synthases (NOS), 404 – NOS isoforms, 405, 406 – regulation of NOS, 406 – splice variants, 406, 407 – subcellular localization, 406 Nitric oxide, 330 NK cells, 445, 446 N-methyl-D-aspartate (NMDA), 105, 107 – receptor, 262, 264, 265, 268, 269, 330, 333, 334, 367, 416–418, 420–422, 424–429, 432–434 No cysteines on the second extracellular domain, 348 Non-MA-ergic neurons, 51, 52 Non-NMDA receptors, 330 Nonsynaptic – interactions, 116 – between neurons, 103 – receptors, 120 Noradrenaline, 23, 103, 104, 107–109 – regulation by 5-HT1A receptors, 175 Noradrenaline (NA) neurons – asynaptic features, 5 – axon morphology, 5 – cell groups, 4 – coeruleocortical NA system, 5 – content of single varicosities, 5 – developmental aspects, 12 – diffuse transmission by, 12 – innervation of – cerebral cortex, 5

– hippocampus, 5 – neostriatum, 4 – spinal cord, 5 – major projections, 4, 5 – morphometric data, 5 – myelencephalospinal NA system, 5, 6 – synaptic features, 5, 6 Noradrenergic neurotransmission – arousal and, 134, 135 – genetics of, 135–140 – stress and, 135 – tonic vs. phasic, 134, 135 Norepinephrine – release, 131, 132, 135 – reuptake, 133–135 – storage, 131 – synthesis, 131, 132 N-P/Q-type Ca2+ Channels, 360, 366 NPPase, 230, 234 NSAIDs, 388, 391, 393, 395 NSD‐1015 (3-hydroxybenzylhydrazine), 154, 155 NTPDase, 230, 234 Nucleoside transporter – concentrative (CNT), 234, 235 – equilibrative (ENT), 230, 234, 235 Nucleus accumbens (NAc), 344, 350, 367, 369 – 5-HT1D receptor transcript location, 181 – 5-HT2A receptor location, 183 – 5-HT4 receptor location, 189 – 5-HT6 receptor location, 194 – 5-HT6 receptor transcript location, 192 Nucleus tractus solitarius – location of 5-HT3 receptor, 187 Obesity, 368, 371 – role of the 5-HT6 receptor, 194 Obsessive compulsive disorder (OCD) – role of SERT, 198 Olfactory bulb – 5-HT5b receptor transcript location, 192 Olfactory tubercle – 5-HT2A receptor location, 183 – 5-HT6 receptor location, 192 – 5-HT6 receptor transcript location, 192 Ondine’s curse, 131 Opioid receptors – regulation of AMPA receptors by, 301 – signaling mechanisms, 301 Outflow nuclei of the basal ganglia, 352

Oxytocin receptors, regulation of AMPA receptors by, 302 P2 receptors – P2X receptors, 228, 235, 236, 238, 239, 243 – P2Y receptors, 228, 235, 237–239, 241, 243 Pain, 232, 243, 245 Panic, links to 5-HT1A receptors, 177 Parkinson’s disease, 269–271, 344, 368 Parkinson’s disease, DA-ergic phenotype expression in, 53, 54 – bienzymatic TH-and AADCexpressing neurons, 59 – monoenzymatic AADCexpressing neurons, 57, 58 – monoenzymatic TH-expressing neurons, 54–57 – origin and properties of striatal neurons, 59–62 PDZ domain-containing proteins, 429, 432, 433 Periaqueductal gray – 5-HT1B receptor location, 179 PGE2 – inhibition of Macrophage Phagocytosis, 388 – mediation of – Ab peptide toxicity, 386, 387, 391–393, 395 – excitotoxicity, 391, 392 – paracrine damage, 395 – phagocytosis, 388 – peripheral functions, 388 – role in ischemic injury and neurodegenerative diseases, 390, 391, 394, 395 PGH2, 386 Phagocytosis, 386, 388, 391–395 Phosphatase, 427, 428 Phosphatidylinositol 3-kinase (PI3K), 416, 432 Phospholipase C – regulation by – metabotropic glutamate receptor, 287 – receptor tyrosine kinases, 304 – serotonin receptors, 298, 299 – tachykinin receptors, 302 Phospholipase C (PLC), 344, 355, 358–359 – activation by 5-HT2A receptors, 183 PI3kinase, 354 Piriform cortex – 5-HT1A receptors, 175 – 5-ht5b receptor transcript location, 192

463

464

Index PLA1, 344, 358, 359 Polymorphism, 344, 351, 352, 363 Pons – location of 5-ht5a receptors, 190 Pore formation, 232, 235, 245 Positively coupled to MAPkinase (MAPK), 354, 355 Positron emission tomography, PET, 76–79 Postsynaptic density, 416, 424 Postsynaptic density protein 95 (PSD–95), 432, 433 Potassium stimulation, 331 Prefrontal cortex, modulation of dopamine release, 175 Presynaptic modulation, 241 Presynaptic neurons, 366 Pretreatment, 83, 87, 89 Primate brain, 76 Prostaglandin (PG) pathway, 386 Protein 4.1 (4.1N), 432 Protein interacting with C-kinase–1 (PICK1), 416, 427 Protein kinase A (PKA), 416, 427 Protein kinase C (PKC), 416, 427 Purinergic, 228, 238, 243, 244, 246 Purkinje cell, 362, 364, 365 Pyramidal neurone – 5-HT1A receptors, 175 – 5-HT4 receptor induced depolarization in hippocampus, 190 – expression of 5-HT2A receptors, 183 – increased activity through 5-HT7 receptors, 194 Radiochemistry, 76, 79, 81, 85, 93, 94 Radioisotope – 11C, 77–81, 84, 89–95 – 13N, 78, 79 – 15O, 78, 79 – 18F, 78, 79, 81, 84, 90, 91, 93–95 Radioligand – carfentanil, 90, 95 – desipramine, 90, 93 – flumazenil, 91, 94 – isoquinoline, 94 – MADAM, 90, 92 – MeNER, 90, 93 – methylspiperone, 91, 92 – raclopride, 89–91 – specific activity, 81 – vinpocetine, 91, 95 – WAY‐100635, 90, 92 Raphe nuclei – SERT location, 196, 197

Receptors – alpha2 (a2), 132 – alphal (a1), 132 – beta (b), 132 – extrasynaptic, 105, 106 – high affinity, 105, 106, 108 – nicotinic, 105, 107 Receptor tyrosine kinases – bFGF receptors, 307, 308 – insulin and IGF receptors, 303, 304 – internalization of AMPA receptors induced by, 303 – neurotrophin receptors, 304 – PDGF receptors, 307, 308 – regulation of AMPA receptors by, 303–308 – role of in synaptic plasticity, 304 – signaling mechanisms, 303 Regulates mesolimbic dopaminergic transmission, 350 Reinforcement, 340, 368, 369 Release, 228–235, 238, 239, 241, 243–246 Release of transmitters, nonvesicular, 106 Retain memory, 350 Retrograde messenger, 365, 367 Retrograde synaptic signaling, 362 Reuptake, of transmitters, 103, 106, 109 RNA editing – 5-HT2C receptors, 186 Schizophrenia, 130, 134–136, 138–140, 351, 368, 371 – role of – 5-HT2A receptors, 184, 185 – 5-HT2C receptors, 186 – 5-HT6 receptors, 193 – 5-HT7 receptors, 196 – SERT, 198 Selective serotonin reuptake inhibitors – action on SERT, 196, 197, 198 – effect on – 5-HT1B receptors, 179 – 5-HT7 receptors, 196 – use with 5-HT1A receptor antagonists, 177 Sensory transmission, 243 Serotonin, 23, 130, 134, 135, 138 – DA synthesis in, 49 – deficiency of, 64 – morphogenetic diffusive factor, 65 – non-MA-ergic neurons, 51–52 – synthetic pathways of, 24 – systemic administration of 5-hydroxytryptophan and, 47

Serotonin (5-HT) neurons – asynaptic features, 6 – cell groups, 6 – content of single varicosities, 7 – developmental aspects, 12 – innervation of – cerebral cortex, 6, 7 – hippocampus, 6, 7 – neostriatum, 6, 7 – spinal cord, 6, 7 – major projections, 6 – morphometric features, 6 – rapheocortical 5-HT system, 6, 7 – rapheospinal 5-HT system, 7 – rapheostriatal 5-HT system, 7 Serotonin receptors, 333 – activation of silent synapses by, 298 – regulation of AMPA receptors by, 299, 300 – signaling mechanisms, 298 SERT – 5-HT1A receptor binding sites and Alzheimer’s Disease, 173 – 5-HT1A receptor, links to depression, 173 – 5-HT2A receptor, links to depression, 173 Signaltransduction, 344, 351, 353–356 Sleep, role of – 5-HT2A receptors, 185 – 5-HT2B receptors, 185 – 5-HT2C receptors, 187 – 5-HT7 receptors, 195 Sodium-dependent transport, 328, 331 Somatostatin receptors – regulation of AMPA receptor by, 301 Specific binding, 82, 83, 85 Sphingomyelin, 355 Spillover – extrasynaptic, 107 – of transmitters, 105, 106 Spinal cord – 5-HT1B receptor location, 179 SR141716A, 350, 355, 361, 365–370 SSRI. See Selective serotonin reuptake inhibitors Stargazin, 430, 432 Stress, 446, 448, 450, 451 – links to 5-HT1A receptors, 177 Striatal neurons, in Parkinson’s disease, 53, 54 – bienzymatic TH-and AADCexpressing neurons, 59 – monoenzymatic AADCexpressing neurons, 57, 58

Index – monoenzymatic TH-expressing neurons, 54–57 – origin, functional properties and significance of, 59–62 Striatum – 5-HT1B receptor transcript location, 179 – 5-HT6 receptor transcript location, 192 Substantia nigra – glutamatergic transmission suppressed by 5-HT1B receptors, 179, 181 – location of – 5-HT4 receptors, 189 – 5-ht5a receptors, 190 – 5-HT6 receptors, 192 – 5-HT7 receptors, 194 – SERT, 196 Superior colliculus – 5-HT1B receptor location, 179 Suprachiasmatic nuclei (SCN), 29 Suprachiasmatic nucleus (SCN) – 5-HT1B receptor location, 179, 181 – 5-HT7 receptor location, 194 Synapse, 416, 420–426, 428–430, 432–434 Synaptic features, 7 Synaptic interaction, between cells, 103 Synaptic plasticity, 262, 264–266, 268, 271, 344, 367, 416, 417, 422, 427, 428, 430, 433, 434 – role of AMPA receptor trafficking in, 284, 285 Synaptic transmission, 261, 262, 264–268 Synaptic vesicles, 327, 331 Synthesis, 228–230, 245 Synthesis of 2-AG, 358, 365, 368 Synthesis-modulating autoreceptors, 162 T helper cells – Th1 cells, 446, 447, 450 – Th2 cells, 446, 447, 450 Tachykinin receptors – regulation of AMPA receptors by, 302 – signaling mechanisms, 288, 298, 302, 303

Taurine, 326–334 – biosynthesis, 327 – deficiency, 327 – distribution, 327 – receptors, 329, 330, 334 – release, 330, 331 – transport, 328, 331 – transporter, 328 Taurine-like immunoreactivity; 327 Thalamus – 5-HT1B receptor transcript location, 178–181 – 5-HT7 receptor location, 194, 196 – SERT location, 196, 198 Therapeutic strategies, 370 TH-expressing neurons, 26, 27, 36 – monoenzymatic, 28, 35 TH-immunoreactive neurons, 35 – distribution of, 46 Three potential N-glycosylation sites, 348 Tolerance, vulnerability, reinforcement, and consumption, 368 Tonic activation, of receptors, 106 Transmission, 114–116, 118–120, 122, 350, 364–368 Transmitter – concentration of, 106–108 – of cytoplasmic origin, 106 – release of, 103–106 Transmitter criteria, 326 Transporter(s), 76, 80, 89–93 – extrasynaptic, 105, 106 – nonsynaptic, 106 Trigeminal ganglia, location of – 5-HT1D receptors, 182 – 5-HT1F receptors, 182 Tuberoinfundibular neurons, 52, 53 Tumors, 452 Tyrosine hydroxylase (TH), 45, 152–154 Tyrosine hydroxylase-immunoreactive neurons, distribution of, 57 Tyrosine kinase, 427, 429 Tyrosine phosphatase, 428 Uptake, 344, 357, 359, 362, 363, 368, 371 Uridine 50 -diphosphate (UDP), 234, 237

Uridine 50 -triphosphate (UTP), 231, 234, 237 Vanilloid receptors, 356 Varicosities – cholinergic, 103, 105 – nonsynaptic, 104 Vasopressin receptors – regulation of AMPA receptors by, 280, 302 Vasopressinergic neurons, tyrosine hydroxylase expression in, 63 Ventral tegmental area (VTA), 344, 366 – 5-ht5a receptor location, 190 – modulation of – dopamine release by 5-HT1A receptors, 175 – noradrenaline release by 5-HT1A receptors, 175 Very high levels in the hippocampus, particularly within the, 352 Vesicular membrane transporter 2 (VMAT2), 25 Vesicular monoamine transporter (VMAT), 156, 158 Vesicular monoamine transporter 2, 45 Virodhamine, 345, 356 VMAT2. See Vesicular monoamine transporter 2 Voluntary alcohol consumption, 369, 370 WIN55212‐2, 356, 365, 366, 369 – g‐Butyrolactone (GBL) and, 162 – innervations in CNS, 165, 166 – life cycle of, 153 – neurotoxins and, 166, 167 – Parkinson’s disease, 151, 154, 167 – receptors, 159–163 – release, 163–165 – rodent MPTP Model, 396 – and schizophrenia, 152, 164 – storage of, 156–158 – structure of, 151 – a-synuclein aggregates, 396 – synthetic pathway of, 152–155 – transporter, 158, 159 – vesicular monoamine transporter (VMAT), 156, 158

465