The Neuromodulators, (International Review of Neurobiology)

International REVIEW OF Neurobiology Volume 64 International REVIEW OF Neurobiology Volume 64 SERIES EDITORS RONALD...

Author: Ronald J. Bradley | R. Adron Harris PhD | Peter Jenner | John Smythies

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International REVIEW OF

Neurobiology Volume 64


Neurobiology Volume 64 SERIES EDITORS RONALD J. BRADLEY Department of Psychiatry, College of Medicine The University of Tennessee Health Science Center Memphis, Tennessee, USA

R. ADRON HARRIS Waggoner Center for Alcohol and Drug Addiction Research The University of Texas at Austin Austin, Texas, USA

PETER JENNER Division of Pharmacology and Therapeutics GKT School of Biomedical Sciences King’s College, London, UK EDITORIAL BOARD PHILIPPE ASCHER TAMAS BARTFAI FLOYD E. BLOOM MATTHEW J. DURING PAUL GREENGARD KINYA KURIYAMA HERBERT Y. MELTZER SALVADOR MONCADA SOLOMON H. SNYDER CHEN-PING WU

ROSS BALDESSARINI COLIN BLAKEMORE DAVID A. BROWN KJELL FUXE SUSAN D. IVERSEN BRUCE S. MCEWEN NOBORU MIZUNO TREVOR W. ROBBINS STEPHEN G. WAXMAN RICHARD J. WYATT


Neurobiology BY

JOHN SMYTHIES Center for Brain and Cognition Department of Psychology University of California San Diego, California, USA and Department of Neuropsychiatry Institute of Neurology London, UK

Elsevier Academic Press 525 B Street, Suite 1900, San Diego, California 92101-4495, USA 84 Theobald’s Road, London WC1X 8RR, UK

This book is printed on acid-free paper. Copyright ß 2005, Elsevier Inc. All Rights Reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopy, recording, or any information storage and retrieval system, without permission in writing from the Publisher. The appearance of the code at the bottom of the first page of a chapter in this book indicates the Publisher’s consent that copies of the chapter may be made for personal or internal use of specific clients. This consent is given on the condition, however, that the copier pay the stated per copy fee through the Copyright Clearance Center, Inc. (www.copyright.com), for copying beyond that permitted by Sections 107 or 108 of the U.S. Copyright Law. This consent does not extend to other kinds of copying, such as copying for general distribution, for advertising or promotional purposes, for creating new collective works, or for resale. Copy fees for pre-2005 chapters are as shown on the title pages. If no fee code appears on the title page, the copy fee is the same as for current chapters. 0074-7742/2005 $35.00 Permissions may be sought directly from Elsevier’s Science & Technology Right Department in Oxford, UK: phone: (þ44) 1865 843830, fax: (þ44) 1865 853333, E-mail: [email protected]. You may also complete your request on-line via the Elsevier homepage (http://elsevier.com), by selecting ‘‘Customer Support’’ and then ‘‘Obtaining Permissions.’’ For all information on all Academic Press publications visit our Web site at www.books.elsevier.com ISBN: 0-12-366865-4 PRINTED IN THE UNITED STATES OF AMERICA 05 06 07 08 09 9 8 7 6 5 4 3 2 1

CONTENTS

Section I.

The Cholinergic System

Part I. Introduction . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. I. Anatomy.. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. Part II. The Muscarinic System. . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. I. Anatomy.. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. II. Electrical Activity. . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. III. Interactions with Other Transmitters. . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. IV. Postsynaptic Cascades . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. V. Other Topics . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. Part III. The Nicotinic System. . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. I. Microanatomy . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. II. Functional Microanatomy . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. III. Interactions with Other Transmitters. . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. IV. Learning . . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. V. Miscellaneous Topics. . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. References . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . ..

Section II. I. II. III. IV. V. VI. VII. VIII. IX.

I. II. III. IV. V.

The Dopamine System

Anatomy.. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. Electrical Reactions. . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. Interaction between Dopamine and other Transmitters . . . . . . . . . . . . . . . . . . .. Dopamine-Induced Signaling Cascades . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. Miscellaneous Items . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. Dopamine and Learning . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. Dopamine and Behavior. . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. Parkinson’s Disease. . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. Miscellaneous Items . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. References . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . ..

Section III.

3 3 4 4 8 19 24 29 55 55 58 61 72 77 80

124 129 132 139 143 148 150 153 155 155

The Norepinephrine System

Microanatomy . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. Functional Neuroanatomy. . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. Electrical Activity. . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. Interactions with Other Transmitters. . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. Postsynaptic Cascades . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. v

174 175 178 180 185

vi VI. VII. VIII.

CONTENTS

Tropins and Cytokines . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . Memory and Learning (LTP) . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . Miscellaneous Items . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . References. . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . .

Section IV.

The Adrenaline System

Text . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . References. . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . .

Section V. I. II. III. IV. V. VI. VII. VIII.

191 192 196 198

213 214

Serotonin System

Anatomy . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . Electrical Actions. . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . Interactions Between Serotonin and Other Neurotransmitter Systems. . . . Postsynaptic Cascades. . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . Serotonin and Synaptic Plasticity . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . Serotonin and the Development of Neural Systems. . . . . . . . . . . . . . . . . . . . . .. . . . Sleep . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . Miscellaneous . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . References. . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . .

Section VI.

218 223 228 236 238 243 244 245 249

Conclusion

Text . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . References. . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . .

269 271

Possibly Obscure Acronyms for Some Readers ........................... Index ........................................................................................

273 275

PREFACE

In my book ‘‘The Dynamic Neuron’’ I gave a comprehensive account of the dynamic process of the growth and pruning of glutamate synapses. In contrast to an older model of the brain, in which its main excitatory synapses were thought to be comparatively long-lasting structures, it is now apparent that these synapses, and the dendritic spines on which most are carried, are continually being removed and regrown in response to activity, or lack of it, at the synapse. Hebbian mechanisms and redox factors play a prominent role in this system. I also described the role of dopamine at the glutamate synapse. The present book extends my coverage to include a comprehensive survey of the role of systems using acetylcholine (both muscarinic and nicotinic), catecholamines (dopamine, noradrenaline and adrenaline) and serotonin in the brain with a particular focus on synaptic plasticity.

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SECTION I THE CHOLINERGIC SYSTEM

Part I. Introduction I. Anatomy A. Macroanatomy Part II. The Muscarinic System I. Anatomy A. Location of Receptor Subtypes B. M1Rs C. M2Rs D. More on Other MRs II. Electrical Activity A. Potassium Currents B. Sodium Currents C. Calcium Currents D. Oscillatory Activity E. Miscellaneous Electrical Topics III. Interactions with Other Transmitters A. ACh and Glutamate B. ACh and GABA C. ACh and Dopamine D. ACh and Interactions with Norepinephrine and Serotonin IV. Postsynaptic Cascades A. MAPKs B. Other Kinases V. Other Topics A. Pathways from MRs to Gene Expression B. Cytoskeleton C. More on Phospholipases D. Adenosine E. Miscellaneous Items F. Reactive Oxygen Species G. Nitric Oxide H. Tropins I. Endocytosis J. Arrestins K. Miscellaneous Items L. Apoptosis M. Neurotoxic EVects of Cholinesterase Inhibitors N. Synaptic Plasticity O. Gene Induction P. Cell Adhesion Molecules Q. Hormones R. Vasopressin and Oxytocin S. T Cells T. Learning

INTERNATIONAL REVIEW OF NEUROBIOLOGY, VOL. 64 DOI: 10.1016/S0074-7742(05)64001-9

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Copyright 2005, Elsevier Inc. All rights reserved. 0074-7742/05 $35.00

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U. Stress V. Behavior W. Sleep X. Alzheimer’s Disease Y. Miscellaneous Part III. The Nicotinic System I. Microanatomy A. Cortex and Hippocampus B. Other Brain Regions II. Functional Microanatomy A. Ionic Conductances III. Interactions with Other Transmitters A. Acetylcholine and Glutamate B. Acetylcholine and GABA C. Acetylcholine and Norepinephrine D. Acetylcholine and Dopamine E. Acetylcholine and Serotonin F. Multiple Interactions: Acetylcholine/Glutamate/GABA G. Synaptic Plasticity H. Postsynaptic Signaling I. Nitric Oxide J. Cyclooxygenase, Arachidonic Acid, and Prostaglandins K. Reactive Oxygen Species L. The ERK/MAPK Pathway M. Endocytosis N. ScaVolding Proteins IV. Learning A. Long-term Potentiation B. The Hebbian Synapse C. Development V. Miscellaneous Topics A. Vasopressin and Oxytocin B. Neurotoxic Anticholinesterases C. Apoptosis D. Tropins and Cytokines E. Melatonin F. Alzheimer’s Disease References

CHOLINERGIC SYSTEM

3

PART I. INTRODUCTION I. Anatomy

A. Macroanatomy The cholinergic system in most of the brain consists of cell bodies located in groups in the basal forebrain and midbrain. These send a sparse and diVuse, but comprehensive, projection to the rest of the brain. The striatum is an exception to this general rule, because most of its cholinergic system consists of local neurons that support a dense local network of cholinergic innervation. The cholinergic input to all areas of the cerebral cortex in primates comes almost exclusively from the nucleus basalis of Meynert (NBM) in the basal forebrain (Bickford et al., 1994). They project to the entire cortical mantle and to local GABAergic interneurons. The cholinergic cells are large with extensive dendritic trees. On these NBM cells acetylcholine (ACh)þ boutons are frequently apposed to spines and small dendrites close to asymmetrical synapses (glutamatergic). There are very few autoreceptors. In the cortex, these NBM neurons have restricted axonal fields that only partially overlap (van der Zee and Luiten, 1999). They terminate mainly on dendritic shafts of both pyramidal cells and interneurons. The basal nucleus of Meynert also projects to the lateral, basolateral, and basomedial amygdala, mainly by GABAergic neurons but with some cholinergic ones as well (Steriade et al., 1987). The nucleus basalis also supplies cholinergic input to certain thalamic nuclei—the intralaminar nuclei (ILN), medial part of the medial dorsal nucleus, the reticular nucleus (Steriade et al., 1990), and to the olfactory bulb. There are, in particular, extensive bidirectional connections between the nucleus basalis and the prefrontal cortex. The main aVerents to the nucleus basalis come from the midbrain pedunculopontine/lateral dorsal tegmental (PP/LDT) nuclei (by a glutamatergic pathway), from the cortex, particularly limbic cortex (Mesulam, 2003), in an ordered topographical array (Za´borszky et al., 1991) and from the intralaminar thalamic nuclei (ILTN), raphe and locus ceruleus (Woolf et al., 1990), as well as a GABAergic input from the amygdala. The second principal cholinergic system is composed of a continuous array of multipolar neurons, with several dendrites directed in all directions, located in the medium septum and diagonal band of Broca (MS/DB). These nuclei project to the hippocampus, medial prefrontal cortex, retrosplenial cortex, and olfactory structures. They supply all cell types on their somata, dendrites, and terminals in all areas of the hippocampus (HC). Unlike NBM cells, MS/DB neurons have many muscarinic receptors on their surfaces. These are (inhibitory) M2Rs on presynaptic local axons and M2Rs and M4Rs on noncholinergic MS/DB cells (van der Zee and Luiten, 1999). The authors suggest that the cholinergic

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innervation of the cholinergic cells in MS/DB, as well as in the few such cells in the HC, may be mainly by volume transmission in which the bouton is en passant and not termineau. In other words, it does not form a specific synapse but releases its transmitter into the intracellular space. Cholinergic interneurons are located mainly in the neocortex, caudate/putamen, nucleus accumbens, amygdala, and olfactory tubercle. The cholinergic nuclei in the midbrain are the pedunculopontine nucleus (PPN) and the lateral dorsal tegmental nucleus (LDT). Their main input includes the nucleus accumbens, hypothalamus, ILN, raphe, and, particularly, the pontine and medullary reticular formation (probably by a glu/asp system). Their main output includes the nucleus basalis, all thalamic nuclei, the amygdala, primary visual cortex (from 20% of LDTN neurons), and primary olfactory cortex (Higo et al., 1996; Sleekier et al., 1994). The hippocampus receives its cholinergic input from the diagonal band of Broca and the medial septum. Minor cholinergic players are the medial habenula, which supplies the interpeduncular nucleus, and the parabigeminal nucleus, which supplies the superior colliculus. PART II. THE MUSCARINIC SYSTEM I. Anatomy

A. Location of Receptor Subtypes Muscarinic receptors occur in all layers in the neocortex but are more common in layers II and V > I, II, and VI > IV. They are also widely distributed in the rest of the brain. There are five types of muscarinic receptor (MR) numbered from 1 to 5. B. M1Rs In the rat brain, levels of M1Rs are high in nucleus accumbens, striatum, hippocampal CA1 neurons and dentate gyrus, and the frontal and parietal cortex, moderate in hippocampal CA2 and CA3 neurons and temporal cortex, and low in cerebellum and spinal cord (Adem et al., 1997). Wei et al. (1994) reported earlier from mRNA studies that M1Rs are located particularly in the rat cerebral cortex. In the primate visual cortex, M1Rs are located in layers 2, 3, and 6 (Tigges et al., 1997). In the rat striatum, Yan et al. (2001) from mRNA studies state that M1Rs (as well as M4Rs) are plentiful on medium spiny neurons (the main projection

CHOLINERGIC SYSTEM

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neurons) of the striatum. In mouse striatum, according to Zhang et al. (2002), the principal cholinergic autoinhibitory receptors are M1 in the striatum (but are M2 in the hippocampus and cerebral cortex). Hersch et al. (1994) show the distribution of muscarinic receptors in the rat striatum. M1Rs in the primate striatum are located postsynaptically on asymmetrical spines (glutamate receptors) on calbindin and medium spiny projection neurons that receive aVerents from the prefrontal cortex (Alcantara et al., 2001). The authors find that M1Rs are also located on symmetrical synapses, where they may modulate inhibitory transmission from local GABAergic neurons and/or from dopaminergic neurons located in the substantia nigra. Species diVerences are important here. For example, in the thalamus in the lateral geniculate nucleus of the rat, M1Rs and M3Rs are located on the somata and dendrites of thalamocortical neurons, whereas M2Rs are presynaptic on interneurons, especially in the reticular nucleus of the thalamus. However, in the cat, M2Rs and M3Rs are located throughout the lateral geniculate nucleus (LGN), and M1Rs are not found (Plummer et al., 1999). C. M2Rs Wei et al. (1994) reported that, in the rat, M2Rs are located particularly in the thalamus, hypothalamus, pons, and medulla. Mrzljak et al. (1998) state that M2Rs in the cortex used to be thought of mainly as autoreceptors on the axons of cholinergic n. basalis cells. They find in the primate cortex that there is, in addition, a widespread localization on noncholinergic neurons and fibers. The M2Rs are found, in the case of pyramidal neurons, only on spines and asymmetrical synapses. On nonpyramidal neurons, they are found on the cytoplasmic surfaces of membranes of their somata, dendrites, and axons. In addition, these workers found that many M2Rs are located perisynaptically and extrasynaptically, suggesting the very important likelihood of cholinergic volume transmission. In rat striatum, M2Rs are located mainly at postsynaptic sites on the somata and dendrites of cholinergic and NPY-somatostatin interneurons and at presynaptic sites on boutons (Bernard et al., 1998). The M2Rs were usually located at extrasynaptic sites and only rarely in association with symmetrical synapses. This constitutes more evidence to suggest that M2Rs mediate volume transmission. However, Iannazzo and Majewski (2000) confirm, in mouse cortex, that presynaptic cholinergic autoreceptors are types M2 and M4 and not M1 or M3. Lesion experiments show that M2Rs are synthesized largely within the cortex and/or are localized mainly at noncholinergic terminals (Mrzljak et al., 1998). In the cat visual cortex, M2Rs are present in all layers except bands in layer 4 (Erisir et al., 2001). In layers 5 and 6 most are presynaptic or are located on axosomatic synapses. In layer 6, some are postsynaptic. These receptors are

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present in both terminals and dendrites in both symmetrical and asymmetrical synapses. In layer 5, 50% of M2Rs are on the dendrites, but very few are on the axon terminals, of GABAergic interneurons. Corticofugal output neurons in the cat brain are particularly modulated by M2Rs (Erisir et al., 2001). In contrast, in rat brain, Tigges et al. (1997) report that M2Rs are located particularly in lower layer 4c and in layer 4a of the visual cortex. Species diVerences may be relevant here (see Plummer et al., 1999). In the cat visual thalamus, there is a dense projection from the parabrachial region to the visual thalamus that modulates sleep/waking and attentional levels (Carden and Bickford, 1999a). There is a dense M2R representation in the neuropil and somata of the LGN, pulvinar, and proximal dendrites of the reticular nucleus. Dendrites of both thalamocortical neurons and interneurons carry M2Rs in both the glomerular and extraglomerular neuropils. These authors state that M2Rs play an important role in blocking thalamic oscillations and in increasing the eYciency of signal transmission during increased attention (Carden and Bickford, 1999a). Likewise, in the lateral posterior nucleus of the cat thalamus (lateral division), there are many M2Rs on the distal dendritic arbors of thalamocortical neurons and of interneurons, as well as on some GABAergic terminals (Carden and Bickford, 1999b). In the cat visual system, acetylcholine modulates visually evoked potentials more in area V1, by a direct cortical eVect, than in the LGN (Arakawa et al., 1997). There is evidence in rats that acetylcholine may modulate visceral neuronal activity in the parvocellular division of the ventral posterior nucleus of the thalamus (Barnabi and Cechetto, 2001). Oda et al. (2001) have studied the cholinergic system in the rat anterior dorsal (AD) and anterior ventral (AV ) nuclei of the thalamus. They found M2Rþ neurons throughout both nuclei. Of these, 30% were on proximal dendritic shafts. A few in AD only were on axon terminals. M3Rs were also seen in both nuclei, mostly on distal dendrites, mainly in the ventrolateral part of the AV nucleus that receives a dense cholinergic input from the dorsal tegmentum. In the rat HC, M2Rs on CA1 neurons are mainly presynaptic on cholinergic and noncholinergic axon terminals. Thus, M2Rs may be heteroreceptors, as well as homoreceptors (Rouse et al., 2000a). M2Rs may inhibit GABA release presynaptically from the terminals of perisomatic inhibitory cells (Hajos et al., 1998), whereas they may stimulate the activity of another class of interneuron that innervates the dendrites of pyramidal cells. Most M2Rþ somata and dendrites are parvalbumin negative, but many terminals colocalize parvalbumin (but not cholecystokinin [CCK] or vasoactive intestinal peptide [VIP]) with the M2R (Hajos et al., 1998). Sugaya et al. (1997) agree that the principal presynaptic autoreceptor in the rat HC is the M2R. But M4Rs are also present in the HC, as evidenced by the presence of their mRNAs, and may also be autoreceptors. M4R RNA is also present on cholinergic neurons in other areas such as the

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medial septum, striatum, reticular formation, and brainstem, as well as on many noncholinergic neurons (Sugaya et al., 1997). In the primate striatum, nearly all cholinergic neurons have M2Rs. Also many noncholinergic NOþ medium aspiny neurons have M2Rs (Smiley et al., 1999). Alcantara et al. (2001) report that M2Rs in the primate striatum are mainly presynaptic on axons of GABAergic neurons. In the striatum, cholinergic interneurons release acetylcholine, which stimulates M1Rs and M2Rs with diVerent eVects on striatal output. Postsynaptic activation of the M1Rs stimulates spiny projection neurons to fire. Stimulation of the presynaptic autoreceptor M2Rs on cholinergic interneurons activates them and so inhibits the firing of the projection neurons (Galarraga et al., 1999). In both rat and primate brain, only 1 to 2% of cortical terminals make asymmetrical contacts onto small dendritic spines of M2R þ cholinergic interneurons in the striatum but, although small, this influence may nevertheless be eVective (Thomas et al., 2000). In the primate nucleus basalis of Meynert, only 10 to 25% of cholinergic neurons have M2Rs (Smiley et al., 1999). There is a special population of NO cells in cortical white matter and adjacent deep cortex that bear M2Rs. These may provide a nitric oxide (NO) relay for input from the nucleus basalis to the cortex (Smiley et al., 1998). Species diVerences also occur in the case of the M2R. For example, Erisir et al. (2001) found that the cat and the monkey have completely diVerent patterns of M2R distribution in layer IV of the visual cortex. D. More on Other MRs Wei et al. (1994) reported that M4Rs are prominent in the rat striatum, and M3Rs and M5Rs have a wide distribution in the brain. In rat striatum, the M4Rs are located mainly in the plasma membrane of somata, dendrite, and spines of medium spiny (projection) neurons. In cholinergic neurons in contrast, the M4R protein is mostly located where it is synthesized, especially in the endoplasmic reticulum (Bernard et al., 1999). Presumably, this is because this protein is destined for export to the axon terminals of these cholinergic neurons. In the primate visual cortex, Tigges et al. (1997) found that M4Rs are located in layers 1, 2, 3, and 5, whereas MR3s are located in layer 4. The cholinergic input to the rat subthalamic nucleus exerts an excitatory postsynaptic eVect mediated by M3Rs (Flores et al., 1996). In the mouse septum, cholinergic septohippocampal neurons are activated by M3Rs but not by M1, M2, or M4Rs (Hsieh et al., 2000). In the rat pituitary, M1Rs and M3Rs are present and active on most cells (Pinter et al., 1999). Stewart et al. (1999) found that mRNAs for MR1, 2, 3, and 4s are present in the rat sensorimotor cortex. In chick retina, M4Rs are predominant during early development, and M2Rs and M3Rs develop later (Nadler et al., 1999).

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A recent study by Yamada et al. (2003) shows that endothelial M(5) receptors mediate vasodilatation in cerebral, but not systemic, arterioles. Furthermore, they showed that M(5) receptors located on dopaminergic nerve terminals play a role in facilitating muscarinic agonist–induced dopamine release in the striatum. The authors state that this is consistent with the observation that the dopaminergic neurons innervating the striatum almost exclusively express the M(5) receptor subtype. The rewarding eVects of morphine and the somatic and aVective components of naloxone-induced morphine withdrawal symptoms are substantially reduced in M5 –/– mice. These behavioral deficits are probably caused by the lack of mesolimbic M(5) receptors, activation of which is known to stimulate dopamine release in the nucleus accumbens (Yamada et al., 2003).

II. Electrical Activity

A. Potassium Currents The brain contains several potassium channels that are modulated by the cholinergic system. There are four main types of Kþ channel. Voltage-activated and Ca2þ-activated channels have six transmembrane domains. ‘‘Leak’’ Kþ channels have four transmembrane domains and inward-rectifying channels* have two. An important group of voltage-activated Kþ channels is the Shaker group (Schrader et al., 2002). This group contains at least five diVerent -subunit proteins derived from the Shaker gene by alternative splicing. Of these, the Shal group (Kv4.1–4.3) participates in the A-type current in neurons, which provides a timing mechanism that regulates the frequency of action potentials and cell excitability. The A-type channel Kv4.2 is found in hippocampal CA1 neuronal dendrites, particularly in the distal branches. Here they play an important role in limiting the peak of back-propagated potentials and in modulating the incoming sensory information (Schrader et al., 2002). Clusters of Kv4.2 channels are also found in neurons of the supraoptic nucleus located under synaptic boutons. Kvbeta is a family of auxiliary proteins that has no transmembrane domain but that impact several features of Kþ channels, including modulating kinetics, exerting a chaperone function, and acting as redox sensors (Schrader et al., 2002). A further group of Kþ channel–associated proteins are the KchIPs, which act as chaperones for Kv4.2 proteins and modulate their kinetics. Phosphorylation of KChIPs may modulate gene transcription down line.

*Inward rectification is a property of channels that conduct inward current at negative membrane potential but little current at positive membrane potentials.

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ATP-sensitive potassium channels are composed of a complex of two structurally diVerent proteins: a member of the inwardly rectifying Kir6 family (Kir6.1 or Kir6.2) and a sulfonylurea receptor (SUR1 or SUR2). The Kir6.1 subunit is predominantly expressed in astrocytes (as well as in a small subset of neurons in the hypothalamic supraoptic and paraventricular nuclei and the striatum) in contrast to the Kir6.2 subunit, which is exclusively expressed in neurons (Thomzig et al., 2003). The voltage-dependent activation of some Kþ channels is further modulated by protein kinases (PKs). Activation of either PKA or PKC shifts the mode of activation to more depolarized membrane potentials (HoVman and Johnston, 1998), whereas activation of the ERK/MAPK system shifts the mode of activation to more hyperpolarized membrane potentials (Yuan et al., 2002). Redox factors also modulate the kinetics of the Kv4.2 channel (Perez-Garcia et al., 1999). The currents these channels conduct include the after-polarization current Iahp (a slow voltage-dependent Ca2þ-activated Kþ current); the M-current IM (a time- and voltage-dependent Ca2þ-independent resting outward rectifying current); a voltage-activated, delayed rectifier-like current Ik; and a leak Kþ conductance (IK leak, which is time-, Ca2þ-, and voltage-independent) (Rouse et al., 2000b; van der Zee and Luiten, 1999). The first two channels induce suppression of firing adaptation (i.e., the inhibition of firing during the course of sustained excitation) that makes the cell more excitable. In CA1 cells in the mouse HC, activation of muscarinic receptors raise the excitability of pyramidal neurons by inhibiting several of these Kþ conductances. In the rat, basolateral amygdala muscarinic activation (probably via M3Rs) inhibits IM, IK, and Iahp potassium conductances but augments the IQ Kþ conductance (which is a hyperpolarization-activated inward rectifying current) (Yajeya et al., 1999). Gabel and Nisenbaum (1999) found that an additional Kþ channel in rat striatum—carrying the 4-aminopyridine-resistant persistent Kþ current Ikrp—can be either suppressed or enhanced by muscarinic receptor activation, depending on the polarization state of the membrane. This is a critical factor in producing the recurring episodes of hyperpolarization and depolarization characteristic of spiny neurons in vivo. In rat lateral geniculate neurons, the activation of MRs leads to a slow depolarization lasting several minutes. This has two components. M1Rs evoke an early large inward current with little change in input resistance. This is due to a decrease in the inward Kþ current IK leak. M3Rs evoke a late small inward current with a large change in input resistance. This is brought about by an increase in the depolarization produced by the cation current Ih (Zhu and Uhlrich, 1998). Cooper et al. (2001) state that, in the brain, the Kþ channel subunit KCNQ2, which is the voltage-activated channel responsible for the M-current, is localized particularly in key areas responsible for the control of rhythmic and synchronized neural activity. From studies on mouse brain, they show that these channels are located on the somata of dopaminergic (DA) and GABAergic cells in the

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substantia nigra; on the large cholinergic aspiny neurons of the striatum; on GABAergic, purinergic, and cholinergic neurons in the septum; on the GABAergic neurons of the reticular nucleus of the thalamus; and on many interneurons, but not pyramidal neurons, in the HC. In the auditory thalamus, acetylcholine modulates parallel lemniscal and nonlemniscal pathways. The former pathway mediates tonotopic relay of acoustic signals, whereas the latter is involved in detecting and transmitting auditory cues of behavioral relevance. Mooney et al. (2004) report that cholinergic muscarinic receptors have opposite membrane eVects on these parallel synaptic pathways. In lemniscal neurons, muscarine induces a sustained membrane depolarization and tonic firing by closing a linear Kþ channel. In contrast, in nonlemniscal neurons, muscarine evokes a membrane hyperpolarization by opening a voltage-independent Kþ channel. The level of membrane hyperpolarization and the strength of local synaptic input determined whether nonlemniscal neurons were suppressed or selectively engaged in detecting and transmitting synchronized synaptic input by firing a high-frequency spike burst. Nonlemniscal neurons predominantly express M2 muscarinic receptors, whereas lemniscal cells had a significantly higher level of M1 receptors (Mooney et al., 2004). GIRKS are G-protein–coupled inwardly rectifying potassium channels that are also modulated by the cholinergic system. In rat brain, these are located densely in the neocortex, thalamus, cerebellar cortex, olfactory bulb, HC, dentate gyrus, and several brainstem nuclei (Ponce et al., 1996). Here they occur postsynaptically on somata and dendrites, as well as on axons and axon terminals. They are especially abundant in the cortical barrels of layer IV of the cortex on thalamocortical terminals. There are two types of GIRK. Luscher et al. (1997) in experiments using GIRK2 knockout mice, have shown that in one type, coded by the GIRK2 gene, the channel is linked to postsynaptic GABA(B), 5-HT1A, or adenosine A1 receptors, activation of any of which evokes an outward current. In the other, muscarinic receptor activation promotes an inward current, which is still present in these mice that lack the GIRK2 gene. In addition, they found that presynaptic receptors for a number of agents, including muscarinic receptors, did not contain the GIRK2 protein. Seeger and Alzheimer (2001) reported an apparent paradox in the muscarinic modulating of somatodendritic processing of EPSPs. In experiments using hippocampal slices, they found that, whereas muscarinic activation increases the excitability of the neuron in several ways, activation of M2Rs on apical dendrites or somata of pyramidal cells reduced the amplitude of EPSPs and glutamateevoked PSPs. This was due to cholinergic activation of a GIRK conductance that served partially to shunt excitatory synaptic input. Some progress has been made in determining the signaling pathways from the AChR to the Kþ channel. These include arachidonic acid (AA) for the GIRK1 subunit in heart tissue (Lohberger et al., 2000) and diacylglycerol (DAG) for M-currents in smooth muscle (Clapp et al., 1992). Recovery from

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the cholinergic inhibition of M-currents requires the production of ATP, lipid phosphorylation by PI4-kinase, and agonist-generated phosphatidylinositol-4,5biphosphate (Suh and Hille, 2002). In the case of the Iahp current, one intermediary is G(q)-activated Ca2þ/calmodulin kinase II (Krause and Pedarzani, 2000; Krause et al., 2002). In cardiac cells, the hydrolysis of phosphatidylinositol biphosphate (PIP2) results in the desensitization of a GTP-binding protein– stimulated Kþ current. Two receptor-regulated pathways cross talk to modulate this current. One pathway signals through the - subunits of G-proteins, which bind directly to the channel and lead to persistent activation of the channel. The second pathway activates PLC, which hydrolyzes PIP2 and limits G-stimulated activity (Kobrinsky et al., 2000). Integrins are cell adhesion molecules found in synapses and elsewhere. GIRK channels bind directly to integrin and require this interaction to maintain the proper localization and function of this channel (McPhee et al., 1998). Stimulation of M3Rs in carcinoma cells activated PKC and leads to an increase in the binding capacity of 1-integrin (Quigley et al., 1998). In the literature of muscarinic receptor research, there is hardly any mention of the scaVolding proteins and other structural proteins that play such a prominent role in ordering the protein machinery contained in the postsynaptic density at the glutamate receptor [see Smythies (2002) for details]. It is, therefore, of interest to come across one report that the interaction of the M2R and its Gprotein (G) with the inwardly rectifying Kþ channel Kir3.2c requires the presence of the scaVolding protein SAP97 (Hibino et al., 2000). The guanylate kinase (K) domain of SAP95 is crucial for this interaction. Acetylcholine-sensitive Kþ channels are directly sensitive to membrane stretch. This eVect is not mediated by G-proteins. This phenomenon is of interest owing to the possible role of membrane stretch in synaptic plasticity (Smythies, 2002). Stretch-sensitive Ca2þ stores have been reported in vascular smooth muscle cells (Mohanty and Li, 2002). These are distinct from the established ryanodine- and NAADP-sensitive stores. B. Sodium Currents Muscarinic receptor activation inhibits the intrinsic bursting electrical activity of rat hippocampal pyramidal cells (Alroy et al., 1999). The pathway for this involves activation of phosphokinase C (PKC), which downregulates the persistent Naþ current carrying slow depolarizing potentials (Mittmann and Alzheimer, 1998). Contrariwise, in guinea-pig lateral septum, stimulation of muscarinic receptors activated a slow after-depolarizing potential (sADP) generated by a cationic conductance largely permeable to Naþ (Carette, 1998). In the neostriatum, activation of M4Rs evoked discharges in a proportion (40%) of neostriatal spiny neurons by activating a Naþ inward current (Figueroa et al.,

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2002). Because the persistent Naþ current (InaP) is activated in the subthreshold voltage range, the cholinergic inhibition of this current would be particularly suited to modulate the electrical behavior of neocortical pyramidal cells at or near their firing threshold (Mittmann and Alzheimer, 1998). In the CA1 area of the HC delayed rectifying Kþ channels have a uniform density, but A-type channels are concentrated in dendrites, where they limit back-propagated potentials, synaptic potentials, and the dendritic initiation of action potentials ( Johnston et al., 2000). C. Calcium Currents Acetylcholine regulates two aspects of calcium activity in neurons. The first is the receptor-mediated entry of calcium into the cell. The second is the redistribution of intracellular calcium in diVerent compartments of the neuron. 1. Receptor-Mediated EVects In the rat striatum, muscarinic receptor activation inhibits Ca2þ currents by two pathways. A PTX-sensitive G-protein pathway inhibits N- and P-currents, whereas a PTX-insensitive pathway inhibits L-currents (Howe and Surmeir, 1995). In the rat striatum, Yan and Surmeier (1996) found that activation of M2Rs and M4Rs on cholinergic interneurons reduces N- and P-type Ca2þ currents by means of a link with Gi/o class G-proteins. In sympathetic neurons, MR activation leads to a slow postsynaptic inhibition of the N-type Ca2þ current at positive potentials and an augmentation at negative potentials. The intermediary signal in the chain mediating this eVect is arachidonic acid (Liu and Rittenhouse, 2003). Presynaptic stimulation-evoked release of acetylcholine in rat striatum is autoinhibited by acetylcholine by both N- and P/Q-type Ca2þ channels (N > P/Q) (Dolezal and Tucek, 1999). In the rat sensorimotor cortex, MRs inhibit Ca2þ currents by two mechanisms (Stewart et al., 1999): a fast voltage–dependent component on N- and P-type currents, and a slow voltage–independent component on L-type currents. Evidence shows that G-proteins are involved in these processes. In gene knockout experiments using sympathetic neurons, Shapiro et al. (2001) found that the fast and voltage-dependent pathway is modulated by M2Rs and the slow and voltage-independent pathway by M1Rs. Neither pathway was aVected in M4R knockout mice. M1Rs also modulated the M-type Kþ current. In snail neurons, activating M receptors promotes the inward current carried by high-voltageactivated Ca2þ channels (Golowasch et al., 1995) by means of ser/thr protein phosphatases. In the entorhinal cortex, muscarinic depolarization is mediated by a Ca2þ channel also largely permeable to Naþ (Klink and Alonso, 1997b). This only

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results in a substantial membrane depolarization if paired with postsynaptic neuronal activity (i.e., a membrane depolarization triggered from some other source that promotes the Ca2þ influx in a voltage-dependent manner. This constitutes a Hebbian mechanism. Gloveli et al. (1999) have studied the eVects of carbachol on the activity of stellate neurons in layer II and pyramidal neurons in layer III in the entorhinal cortex. Carbachol induces a membrane depolarization associated with synaptic oscillations in both types of neuron. In contrast to the intrinsic theta rhythm of stellate cells with one peak at 7 Hz, muscarinic activation produced mixed and frequencies with peaks at 11, 23, and 40 Hz associated with a rise in intracellular Ca2þ levels. In the stellate cells in layer II, low-voltage-activated Ca2þ channels are predominant, whereas in layer III it is high-voltage Ca2þ channels that are dominant in this response (Gloveli et al., 1999). In a previous study, this group (Gloveli et al., 1997) determined that, in the rat entorhinal cortex, short train repetitive stimulation below the intensity required to generate action potentials induced a slow hyperpolarization that had three components. The first lasting 1 second, was cholinergic. The second, lasting 6 seconds, was GABAergic, and the third (20 seconds) was opioid. In rat subiculum, MR activation inhibits burst-afterhyperpolarizations and discloses long-lasting depolarizing plateau potentials that are mainly the result of a Ca2þ-dependent nonselective cationic conductance (Kawasaki et al., 1999). The rise in intracellular calcium levels activates guanyl cyclase, and the cGMP produced activates cyclic nucleotide-gated channels, which induce the plateau potential (Kuzmiski and Mac Vicar, 2001). These potentials powerfully modulate the intrinsic excitability of subicular bursting cells. In the CA1 cells of the HC, plateau potentials generated by M1/M3Rs require protein dephosphorylation induced by activation of a protein phosphatase (Fraser et al., 2001). Qian and Saggau (1997) report that, in rat hippocampal slices, the inhibition of synaptic transmission at the CA3–CA1 synapse by MRs is largely due to inhibition of presynaptic voltage-dependent calcium channels. In studies on hippocampal slices, cholinergic stimulation by local microapplication augments CA1 firing by two routes. On somata, acetylcholine suppresses the slow afterhyperpolarizations that follow repetitive stimulation. On dendrites, acetylcholine augments slow tetanic depolarization by directly up-regulating NMDA currents, thus potentiating Ca2þ signaling (Egorov et al., 1999). The authors suggest that these data show that subcellular local muscarinic modulation may contribute to parallel information processing and storage by the dendritic synapses of CA1 neurons. Neuromodulation by transmitters in the central nervous system usually aVects cell excitability by modifying the membrane potential, discharge patterns, and spike frequency. A new way to aVect this has been described by Figenschou et al. (1996)—by changing the waveform of the action potential. In CA1 cells in the rat

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HC, low doses of carbachol caused an increase in spike duration. High doses or long-lasting applications also reduced the spike amplitude and rate of rise and raised the spike threshold. The intermediary mechanism is Ca2þ dependent and probably involves PKC. In further experiments on hippocampal slices, Fraser and Mac Vicar (1996) confirmed that cholinergic stimulation results in a long-lasting plateau potential, as well as a slow aftepolarization. They conclude that the plateau potential represents a novel regenerative event representing the interplay between highvoltage activated Ca2þ channels and Ca2þ-activated nonselective cation conductance. The slow afterpolarization represents activation of the latter conductance by residual intracellular Ca2þ. Wheldon et al. (2001) showed in neuroblastoma cells that M3R activation increases intracellular Ca2þ by an inositol 1,4,5-triphosphate–dependent mechanism. In HEK cells and in the exocrine avian nasal gland, Ca2þ entry leading to oscillatory Ca2þ signals depends not on inositol phosphates but on the PLA2mediated generation of arachidonic acid (Shuttleworth, 1996; Shuttleworth and Thompson, 1998). In the rat neocortical pyramidal cells, activation of MRs raises cell excitability by releasing Ca2þ from inositol 1,4,5-triphosphate (IP[3])–sensitive intracellular Ca2þ stores. This function is not present in 1-weekold rats who possess functional IP(3)Rs and Ca2þ channels, but these are not yet functionally connected (Yamamoto et al., 2002a,b). Naþ-dependent action potentials, produced when the neuron fires, backpropagate over the dendritic tree. This process plays a key role in Hebbian synapses (Smythies, 2002). These potentials normally decrease, so that later spikes in the chain decrease in the dendritic arbor. In mouse HC, both local depolarization and MR activation inhibit this incremental decrease, so that all spikes in the back-propagated activity are of the same amplitude when recorded from proximal and middle (but not apical) dendrites (Tsubokawa et al., 2000). Back-propagated potentials increase intracellular Ca2þ levels in all dendritic locations (Tsubokawa and Ross, 1997) but by diVerent intracellular mechanisms in the case of local depolarizations (by means of CaMKII) and after MR activation (Tsubokawa et al., 2000). So the cholinergic system may induce a Hebbian eVect at the synapse by facilitating back-propagation and so prolong the period of depolarization in the postsynaptic membrane simultaneous with the incoming volley needed for the Hebbian eVect. In the hippocampal CA1 pyramidal neurons, these back-propagated potentials are potentiated either by -adrenergic stimulation (by means of PKA) or by muscarinic stimulation (by means of PKC) (HoVman and Johnston, 1999a, b). In guinea-pig myocytes, -adrenergic agonists stimulate Ca2þ currents by way of activation of adenylyl cyclase and the synthesis of cAMP. This eVect was at first inhibited by MR activation with a following transient stimulatory rebound. The stimulatory action is due to a direct activation of adenylyl cyclase

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by the subunits of the cholinergic-receptor bound G protein (Belevych et al., 2001). Growth hormone–releasing hormone (GHRH) induced Ca2þ-mediated oscillations in membrane potential in rat somatomorphs grown in culture. These oscillations were blocked by cholinergic agonists and somatostatin (Kwiecien et al., 1997). In the CA1 region of the HC, whereas Na2þ channels have a uniform distribution, calcium channels of the L- and N-class channels are concentrated on the soma and R- and T-class channels are concentrated on dendrites ( Johnston et al., 2000). 2. Redistribution of Intracellular Calcium Cholinergic receptor activation evokes complex changes in the distribution of intracellular calcium (Beier and Barish, 2000). In the CA1 pyramidal cells of the mouse HC, muscarinic stimulation results in enhanced accumulation of Ca2þ in the cytosol, presumably by release from stores, in the absence of action potentialdriven Ca2þ entry. By modulating the time course of Ca2þ signals, MR activation may regulate the activation of Ca2þ-dependent intracellular processes. Muscarinic receptors also modulate calcium waves in neurons. In CA1 hippocampal neurons, Power and Sah (2002) report that serial activation of MRs evokes a focal rise of free calcium in the apical dendrite that propagates as a wave down to the soma and invades the nucleus. Inositol 1,4,5-triphosphate receptors are essential for this response, which eVects the transcription of novel genes. D. Oscillatory Activity 1. Hippocampus MR activation in the hippocampus induces oscillatory responses whose frequency is nonlinearly dose-dependent. In the rat HC, low doses induce asynchronized , medium doses synchronized , and high doses induce synchronized (Fellous and Sejnowski, 2000; Tiesinga et al., 2001). The generation of hippocampal depends on complex interactions between MRs and mGluRs (Cobb et al., 2000). An I(h) pacemaker current located in the CA3 region is important for the generation of this activity (Cobb et al., 2003). M1Rs are necessary for oscillations in mouse hippocampal CA1 neurons. M1R activation depolarizes CA3 pyramidal neurons by increasing the mixed Naþ/Kþ current I(h) and by increasing the Ca2þ-dependent nonspecific cation current I(cat) and not by modulating I(M). Local activity resulting from MR activation in the CA1 area of rat hippocampus may be produced by CaMKIIinduced suppression of the slow I(AHP) and the induction of plateau potentials

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(Alberi et al., 2000). In rat HC, interneurons near the border between stratum radiatum and stratum lacunosum-moleculare (LM) display intrinsic membrane potential oscillations at the frequency when depolarized near threshold. MR activation depolarized these cells and induced . This may contribute to the generation of rhythmic inhibition that paces intrinsically generated in CA1 pyramidal cells (Chapman and Lacaille, 1999). However, Williams and Kauer (1997) caution that MR-induced oscillations are fundamentally diVerent from the in vivo rhythm. In their experiments, they showed that carbachol produced a regularly spaced bursting pattern of field potentials in both the CA1 and CA3 regions of rat HC. The oscillatory generator was localized in CA3. These oscillations required AMPAR activation, but the spread of activity to CA1 required NMDAR activation. The authors state that their data are more consistent with epileptiform bursting than rhythm. They suggest that the fundamental oscillator for this activity is a slow rhythmic oscillation induced in CA3 cells by carbachol that excites multiply connected CA3 neurons. The generation of this activity depends on M1Rs, M3Rs, and nicotinic AchRs, but not on GABA nor mGluRs. The hippocampal rhythm is driven from the medium septum and diagonal band (MS/DB) by a phasic inhibition of cholinergic MS/DB neurons by GABAergic MS/DB neurons followed by a rebound of their firing (Brazhnik and Fox, 1997). This leads to the recruitment of the whole MS/DB neuronal population into the synchronized rhythmic bursting pattern that drives hippocampal . Low-frequency oscillations (2–7 Hz) in the rat subiculum rely on a MR-dependent activation of an intrinsic I(CAN)-dependent oscillatory mechanism and are integrated within the network by means of non-NMDA receptor-mediated transmission (D’Antuono et al., 2001). In rodent thalamic brain slices, MR activation depresses low-frequency responses in corticothalamic synapses more than it does high-frequency responses. In whole brain, stimulation of the brainstem reticular formation strongly inhibits low-frequency but not high-frequency corticothalamic responses. Thus, during muscarinic-evoked arousal, only high-frequency inputs from the neocortex are allowed to reach the thalamus (Castro-Alamancos and Calcagnotto, 2001). In the lateral posterior and lateral geniculate nuclei of the thalamus, activation of MRs switch the firing pattern in thalamic interneurons from bursting to tonic by selectively decreasing their input resistance rather than by altering their membrane potential as in the case of some other neuromodulators (Zhu and Heggelund, 2001). This is due to the muscarinic enhancement of a hyperpolarizing Kþ conductance and two depolarizing cation conductances. This action electronically uncouples the distal dendrites of thalamic interneurons and mediates the cholinergic transformation of long-range, slow dendrite-, and axon-mediated inhibition into short-range, fast dendrite–originated inhibition.

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Thus MRs dynamically modulate the electronic properties of the dendrites of thalamic interneurons in this way. Dopamine strongly depresses cholinergic oscillations in area CA3 of rat hippocampus by activation of D1-like dopamine receptors, probably by means of impairment of interneurons involved in generation and maintenance of the carbachol-induced network rhythm (Weiss et al., 2003). 2. Cortex In the guinea-pig entorhinal cortex, MR activation induces synchronous activity of an epileptiform nature driven by cortically projecting layer V neurons. Layer II (but not layer III) neurons can also generate MR-activated oscillatory activity. This epileptiform activity required activation of AMPA but not NMDA receptors (Dickson and Alonso, 1997). In this brain area, fast oscillations induced by MR activation are mediated by direct muscarinic excitation of a highly localized reciprocal excitatory–inhibitory network in the superficial layers of the medial entorhinal cortex (Dickson et al., 2000). The entorhinal cortex contains both stellate cells (SC) and nonstellate cells (non-SC). MR activation plays an important role in tuning their oscillatory dynamics by causing diVerent types of membrane depolarization in each (Klink and Alonso, 1997a). In SCs, MR activation depolarization was associated with subthreshold membrane potential oscillations and clusters of spike discharges. In non-SCs, MR activation drastically aVected firing behavior by promoting the development of voltage-dependent, long-duration, slow bursts of action potentials. Concomitantly, the slow afterhyperpolarization was replaced by long-lasting plateau postdepolarizations. In both, MR activation produced conspicuous changes in the action potential waveform and its afterpotentials of a nature to suggest modulation of a voltage-dependent Naþ conductance. This suggests that the entorhinal cortex supports two parallel processing systems that could diVerentially aVect their hippocampal targets (Klink and Alonso, 1997a). In mouse prefrontal cortex, M2Rs are autoreceptors for acetylcholine release. This activates the EEG in the contralateral cortex (Douglas et al., 2002). In the rat neocortex, requires simultaneous MR and GABA(A) activation, as well as an intact glutaminergic transmission (Lukatch and Mac Iver, 1997). The role of acetylcholine during arousal is more than simply biasing cortical neurons toward greater excitability. Rather, Dancause et al. (2001) showed in experiments using rats that MRs on inhibitory interneurons, or on the dendritic terminals of pyramidal cells, in the superficial layers of the cortex enhance the first intracortical synaptic events in the somatosensory cortex but reduce the population response at later times during the first 250 ms after a tactile stimulus.

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Cortical arousal is associated with low-voltage fast activity (LVFA). This is mediated by two distinct inputs to the cortex. LVFA produced by stimulation of the amygdala and locus ceruleus depends on cholinergic mechanisms by way of the nucleus basalis of Meynert. Here cells that promote LVFA are stimulated, whereas other cells that promote cortical slow activity are inhibited. LVFA produced by stimulation of the dorsal raphe and the superior colliculus are mediated by serotonin. Stimulation of the orbitofrontal cortex activates both pathways (Dringenberg and Vanderwolf, 1997). In human somatosensory cortex, stimulation-induced high-frequency oscillation generated by specialized pyramidal cells (‘‘chattering cells’’) is significantly enhanced by cholinergic activation. In contrast, acetylcholine does not modify the firing rate of fast-spiking GABAergic interneurons (Restuccia et al., 2003). 3. Other Areas Blockade of presynaptic M2Rs and activation of postsynaptic M1Rs in the reticular nucleus of the rat thalamus inhibit thalamocortical oscillations and evoke neocortical arousal (Puolivali et al., 1998). Cells in the anterior pituitary cell line GH3 have MRs that excite Kþ channels and inhibit Ca2þ channels, leading to a decrease in cell excitability and to a reduction in the frequency of Ca2þ oscillations by 85% (Kushmerick et al., 1999). Striatal cholinergic interneurons are endogenously active and generate action potentials in the absence of any synaptic input (Bennett and Wilson, 1999). In a similar manner, cortical inhibitory GABAergic neurons have their own cholinergic-dependent mechanism for generating slow rhythms (0.1–0.3 Hz) independent of excitatory circuits. E. Miscellaneous Electrical Topics In the rat HC using whole-cell recordings, McQuiston and Madison (1999a) found that there are four types of GABAergic interneurons with respect to their reaction to acetylcholine. In type 1 found in all layers of CA1, acetylcholine induces a depolarization by two mechanisms (1) inhibition of a Kþ current and (2) an eVect depending on extracellular Naþ. In type 2, the response was a hyperpolarization and in type 3 a biphasic eVect of a hyperpolarization followed by a depolarization. Type 4 neurons showed no reaction to acetylcholine. In a further series of experiments, McQuiston and Madison (1999b) found that most CA1 neurons react to direct stimulation by injected current or synaptic stimulation with a depolarization followed by an afterhyperpolarization (AHP). MR activation converted this AHP to an afterdepolarization (ADP) often of such magnitude as to evoke action potentials.

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Activation of M1Rs in the deep entorhinal cortex leads to a powerful inhibition of field potentials in layer II of the entorhinal cortex. This is not mediated by suppression of presynaptic Ca2þ signals. This activity should be important for the shaping of oscillatory activity and synaptic plasticity during cholinergic-induced oscillations (Richter et al., 1999).

III. Interactions with Other Transmitters

A. ACh and Glutamate 1. Presynaptic Interactions In rat hippocampal CA1 cells, stimulation of input glutamatergic SchaVer collaterals produces EPSCs (De Sevilla et al., 2002). Simultaneous activation of MRs selectively inhibits glutamate release at functional synapses but not at silent synapses. This eVect is produced by non-NMDARs. The authors conclude that the SchaVer collaterals have two types of terminals, one in functional synapses selectively sensitive to regulation by MRs and the other at silent synapses less sensitive to that regulation. Thus, the specific inhibition of functional synapses would have the significant eVect relating to synaptic plasticity of favoring activity mediated by NMDARs at silent synapses without the potentially disrupting modulation of non-NMDARs and functional synapses. Cholinergic presynaptic inhibition of neostriatal glutamatergic transmission is mediated by means of Q-type calcium channels (Barral et al., 1999). M3Rs located at a presynaptic site on glutamate axonal terminals on dopaminergic cells in the mesencephalon inhibit glutamate release and excitatory synaptic potentials (Grillner et al., 1999). In experiments using brain slices, acetylcholine exerted two diVerent eVects on the glutamatergic fast-spiking projection neurons (Koos and Tepper, 2002). On the synapses that fast-spiking GABAergic interneurons form on the glutamatergic projection neurons, MRs inhibited the release of GABA presynaptically, leading to a disinhibition of the projection neuron. Simultaneously, acetylcholine directly activated the GABAergic interneurons by means of nicotinic receptors, thus balancing the first eVect. Corticostriatal projection neurons release glutamate in mouse striatum, producing negative field potentials by means of activation of AMPARs (Niittykoski et al., 1999). Activation of presynaptic M3Rs (but not M2Rs) inhibited this eVect. This same release is also inhibited by opioid receptors (Rawls et al., 1999). Activating MRs in rat auditory cortex first induces a non–G-protein linked reduction in the amplitude of glutamate-induced (NMDARs) depolarizations in

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layer II to III pyramidal neurons, followed by a long-lasting G-protein–induced enhancement (Aramakis et al., 1997b). In the reverse direction in cultured rat septal neurons, Suzuki et al. (2003) showed that endogenously released glutamate produced ACh release by stimulating AMPA receptors. Fournier et al. (2004) report that glutamate can activate cholinergic basal forebrain neurons by means of both AMPA and NMDA inotropic receptors but has a more modest eVect on EEG activation. 2. Postsynaptic Interactions Blockade of NMDARs can lead to neurotoxicity. Farber et al. (2002) suggest this eVect is in part mediated by the blockade of NMDARS on inhibitory neurons in multiple subcortical brain regions. This would lead to a disinhibition of glutamatergic and cholinergic projections to the cortex and thus excess neurotoxic excitation of cortical neurons. Likewise, in the cerebellum, M2R activation inhibits GABA-releasing Golgi cells and so promotes glutamate release from parallel fibers ( Johnson et al., 2000). In hippocampal CA1 cells, stimulation of MRs activates PKC and its downstream Src signaling system. This phosphorylates the NMDAR and upregulates NMDA-evoked currents (Lu et al., 1999). Wozniak et al. (1998) also found that MR activation increased the neurotoxic eVect of NMDAR blockade. The principal feature of this neurotoxicity was degeneration of synaptic spines and loss of synaptic complexes—an important finding we will return to later. On the other hand, in cultures of hippocampal cells, blockade of muscarinic receptors by scopolamine protected against the neurotoxicity induced by glutamate (Rami et al., 1997). Glutamate, by means of inhibitory type 2 metabotropic receptors on cholinergic interneurons, also inhibits the release of acetylcholine from these neurons (Pisani et al., 2002). These authors suggest that this eVect is mediated by P-type high-voltage–activated Ca2þ channels, a lowering of intracellular calcium levels, and inhibition of plateau potentials. Vogt and Regehr (2001) have studied the cholinergic modulation of excitatory (glutamatergic) synaptic transmission mediated by associational/commissural (A/C) fibers and mossy fibers (MF) in the CA3 region of the rat hippocampus. Their results were complex. At the A/C synapse, MR stimulation inhibited the synapse directly by inhibition of presynaptic calcium channels, thus downregulating glutamate release. At MF synapses, muscarine elevated the firing rate of inhibitory cells, which increased GABA release and so inhibited MF receptor activation indirectly. Nicotinic receptors could also be involved in the excitation of inhibitory neurons ( Vogt and Regehr, 2001). In cultured striatal neurons, activation of both MRs and MNDARs leads to the release of arachidonic acid (AA). Tence et al. (1995) report that there is a strong synergistic eVect if these receptors are stimulated at the same time. This

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eVect depends on the massive inflow of Ca2þ induced by NMDAR stimulation and the coupling of the MR with PLA2. B. ACh and GABA 1. Presynaptic and Postsynaptic Interactions Acetylcholine can augment or inhibit GABAergic activity depending on circumstances. In rat auditory cortex, MR activation inhibits both the fast IPSPs mediated by GABA(A) and the slow IPSPs mediated by GABA(B). This leads to disinhibition of glutamatergic fast EPSPs. MR activation modulates fast glutamatergic neurotransmission diVerentially as a function of the strength of inhibition, so that activity produced by ‘‘weak’’ inputs is suppressed and that evoked by ‘‘strong’’ inputs is enhanced (Aramakis et al., 1997a). Also in rat auditory cortex, repetitive stimulation at low stimulus intensities gives rise to a hyperpolarizing response, and stimulation at a higher frequency elicits a depolarizing response. The former is mediated by MRs on GABAergic interneurons. The latter depends on activity at AMPARs, NMDAs, and MRs (Bandrowski et al., 2001). Using microinjection techniques in rat visual cortex, Kimura and Baughman (1997) showed that, in addition to the well-established fact that activation of M3Rs directly facilitates EPSPs, activation of M1Rs inhibits EPSPs by a presynaptic mechanism and activation of M4Rs inhibits IPSPs also by a presynaptic mechanism. M3Rs also excite GABA INs in the developing rat inferior colliculus (Yigit et al., 2003). In rat MSDB, AChE stimulates most GABA neurons (excess ACh being released from recurrent collaterals of the cholinergic septohippocampal neurons) by way of muscarinic (mainly the M3R), but not nicotinic, receptors (Wu et al., 2003). Kawaguchi (1997) has studied the diVerential eVects of muscarinic agonists in rat frontal cortex on the GABAergic interneurons containing diVerent polypeptides. He found that neurons containing somatostatin or VIP were depolarized with spike firing. Large cells containing CCK were hyperpolarized followed by a slow depolarization. Smaller CCK cells were only depolarized. Cells containing the protein parvalbumin were not aVected. In rat striatal slices, Harsing and Zigmond (1998) have described complex interactions among acetylcholine, GABA, and dopamine. GABA release is stimulated by M1Rs and is inhibited by M3Rs. Dopamine inhibits GABA release by means of its D2 receptors. The M1Rs and D2Rs may be located postsynaptically on medium-sized spiny GABAergic projection neurons. The thalamus receives a dense cholinergic input from the PPN that regulates thalamic function during waking. One mechanism concerned is the inhibition,

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in rat thalamic slices, of GABA release by M2Rs, located on the dendrites of GABAergic Ins, as well as on the somata and proximal dendrites of GABAergic cells in the thalamic reticular nucleus (Rowell et al., 2003). The GABA transporter GAT1 is endogenously expressed in rat hippocampal neurons. MR stimulation inhibits GAT1 function by means of PKC by promoting its endocytosis (Beckman et al., 1999). C. ACh and Dopamine 1. Basics The striatum is concerned in sensorimotor planning and learning. The interactions between dopamine and ACh play a major role in this activity. Only 2% of striatal neurons are cholinergic, but these provide a dense local innervation. They fire tonically at 5 Hz, which promotes dopamine release. During a conditioned motor task, the cholinergic interneurons stop firing, which inhibits dopamine release. Cholinergic neurons in the PPN and the LDTN in the midbrain and pons project to dopaminergic neurons in the ventral tegmental area. Here they activate DA neurons quickly (by way of nicotinic receptors) and also activate them slowly by way of muscarinic receptors (Yeomans et al., 2001). The former eVect is related to the nicotine ‘‘high.’’ The latter eVect (mediated by M5Rs) is needed for the rewarding eVects of hypothalamic stimulation. 2. Presynaptic Interactions The complex interactions between GABAergic and cholinergic neurons acting on dopamine release in the striatum of freely moving rats are reported by Whitehead et al. (2001). Dopamine release is modulated by both cholinergic and GABAergic tonic inhibitory mechanism by way of MRs and GABA(A)Rs, respectively. In addition, NMDA-stimulated DA release involves obligatory muscarinic facilitation and an inhibitory component from the GABA(A) system. So, as is often the case, the cholinergic system can promote or reduce an eVect, depending on circumstances. In the past, ACh was thought simply to promote Kþ-stimulated DA release in the striatum. Zhang et al. (2002) have shown that things are not that simple. In striatal slices from M1–M5 knockout mice, they show that each type of muscarinic receptor has its own specific eVect. M4Rs located on cell bodies promote DA release. M3Rs on axon terminals inhibit DA release (by way of promoting striatal GABA release). M5Rs also on axon terminals promote DA release (by inhibiting striatal GABA release). No function could be ascribed in this tissue to M1 or M2 receptors.

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3. Postsynaptic Interactions In DA neurons in rat mesencephalic slices, muscarinic ACh M3Rs, by means of a G-protein, reduce EPSCs evoked by local electrical stimulation. The mechanism involved is not known (Grillner et al., 1999), except that it is not due to an eVect on N-, P-, or Q-Ca2þ channels. In rat VTA, MRs activate DA cells, leading to the release of DA in the VTA and in its projection areas (Gronier et al., 2000). According to Ince et al. (1997), 95% of rat nigrostriatal neurons bear D1Rs, and 92% bear M4s: only 1% have D2Rs. This suggests that D1Rs and M4Rs are colocalized, whereas D1Rs and D2Rs are segregated. In contrast, nearly all striatal interneurons contain D2 and D5, but no D3 or D4, mRNAs (Yan et al., 1997). These workers found that activation of D2 DA receptors on cholinergic interneurons reduces N-type Ca2þ currents by means of a membrane-delimited Gi/o class G-protein that is not regulated by PKC. They suggest that this mechanism may underlie the ability of D2Rs to reduce striatal ACh release. In rat striatum, stimulation of M4Rs inhibited the activation of adenylyl cyclase induced by dopamine (D1) (DeLapp et al., 1996; Olianas et al., 1998). This eVect was also found in rat nucleus accumbens (Onali and Olianas, 2002). This provides evidence that M4Rs and dopamine D1 receptors are colocalized and functionally interlinked. Also in rat striatum, activation of both D2Rs and of MRs inhibited excitatory synaptic transmission by separate nonadditive and, indeed, antagonist mechanisms. Thus, the eVect of MR stimulation depends on the extent of D2R activation, suggesting cross talk between these two systems at this level. The eVect may be due to a shared PKA-dependent messenger system (Hsu et al., 1996). Pisani et al. (2000) describe the complex interaction between GABAergic, cholinergic, dopaminergic, and glutamatergic systems in the striatum. Cholinergic interneurons have four postsynaptic inputs—an excitatory glutamatergic input from the cortex, an excitatory dopaminergic input (D1) from the substantia nigra, as well as inhibitory (local) GABAergic and cholinergic inputs. The DA system also has inhibitory synapses (D2) on the terminals of the GABAergic and cholinergic axons. Stimulation of cholinergic receptors can also induce a very important Hebbian eVect. Cheramy et al. (1996) found that such activation stimulates PKC. This, in turn, phosphorylates the NMDA receptor so as to induce a conformational change, which removes the Mgþþ ion that normally blocks the ion channel linked to this receptor. As we will see, cholinergic mechanisms can induce Hebbian eVects by other means. Tsukada et al. (2000) have presented the novel hypothesis that cholinergic mechanisms modulate DA turnover in the striatum by changing the dynamics of DA synthesis and of the availability of the DA transporter DAT. Varoqui and Erickson (1997) give a general review of vesicular neurotransporters.

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Activation of D2 dopamine receptors can protect against excitotoxicity resulting from excess activity at both glutamate and cholinergic receptors (Bozzi and Borrelli, 2002). In the rat nucleus accumbens, CCK-A inhibits GABA release acting by means of local M2Rs (Ferraro et al., 1996). In this system, CCK-B promotes GABA release. Neurons in various regions of the brain are activated by CCK. Activation of muscarinic receptors can decrease or even reverse this eVect (Davidowa et al., 1998). In rat striatum, substance P activates cholinergic neurons in striatal slices, resulting in increased DA outflow (Khan et al., 1996). Later work by the same group (Khan et al., 2000a) showed that the eVect of substance P on DA outflow in this preparation depends on the balance between its eVects on M1Rs (inhibitory) and M2Rs (excitatory). Dopaminergic neurons in the rat ventral tegmental area (VTA) project to the nucleus accumbens, and their release of DA is under tonic GABAergic inhibition and tonic cholinergic excitation mediated by both MRs and NRs. The dopaminergic neurons in the VTA may be under feedback inhibition mediated by the nucleus accumbens GABA system (Rahman and McBride, 2002). D. ACh and Interactions with Norepinephrine and Serotonin See under these latter sections.

IV. Postsynaptic Cascades

A. MAPKs The second messengers preferentially involved in muscarinic activity are cAMP for M2Rs and M4Rs and the PLC–phosphoinositol/Ca2þ pathway for M1R, MR3, and MR5 receptors (plus in some cases cAMP) (Murthy and Makhlouf, 1997), but there is some overlap. The muscarinic receptors activate a number of postsynaptic cascades, including the mitogen-activated protein ser/ thr kinase (MAPK) pathway that induces the phosphorylation of key protein molecules in the neuron often by means of a second protein kinase leading in many cases to changes inter alia in ion channel activity. The activated MAPK can also traYc to the nucleus, where it can aVect gene expression and protein synthesis. In the cortex and hippocampus, the dominant muscarinic receptor is M1. However, in this location in mouse forebrain, all five muscarinic subtypes can activate PLC (Hamilton and Nathanson, 2001). In PC12 cells, whereas M1Rs represent only a small proportion of MRs in the cell (most are M4Rs),

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nevertheless they are responsible for activating the chain PKC ! MAPK ! protein phosphorylation (Berkeley and Levey, 2000). Likewise, in mouse hippocampal slices, it is the M1R that is responsible for activation of extracellular signal–regulated kinase (ERK—a type of MAPK) (Berkeley et al., 2001). In the pancreatic acinar cells, both M1Rs and M3Rs (but not M2Rs) stimulated MAPK activity, leading to expression of the early response genes c-fos and c-jun (Turner et al., 2001). In contrast, in transfected CHO cells, M3Rs activate both the ERK and another pathway the c-jun N-terminal kinase ( JNK), but the M2R activates only the former (Wylie et al., 1999). In oligodendrocytes, MRs activate MAPK by way of the PKC pathway leading to c-fos expression and cellular proliferation (Larocca and Almazan, 1997). In rat neural precursor cells, MRs play an important role in neurogenesis. This is mediated by the cascade ! increased intracellular (i.c.) Ca2þ levels ! activation of MAPK ! increased DNA synthesis and increased production of MAP2(þ) (a neuronal specific marker) in which processes the nonreceptor protein tyrosine kinase c-Src and CREB signaling are essential (Zhao et al., 2003). These MAPK pathways start with the MR attached to a G-protein. DiVerent systems can use diVerent G-proteins. In mouse hippocampal CA1 pyramidal cell neurons, the MR activating ERK is coupled to a G(q) G-protein. In this tissue, stimulation of mGluRs also activates ERK in glia—this time coupled to a G(i/o) G-protein (Berkeley and Levey, 2003). In HEK293T cells, both M1Rs and M3Rs stimulate ERK using the simultaneous and synergistic employment of two parallel pathways, one based on the G-protein G(q), leading to PKC activation. The other is based on G (i), leading to Ras activation (G/ subunits, PI3-K, and tyrosine kinases are not involved). This stimulation leads to the transcription of Elk1 (Blaukat et al., 2000). In kidney cells, similar parallel pathways exist using G(q) and G/ G-proteins (Yamauchi et al., 2001). Signaling to a new form of MAPK—ERK5—involves the G-proteins G(q) and G(12/13) and not G(i), G(s), or G subunits (Fukuhara et al., 2000). This variability is marked in pancreatic cells, where M2Rs and M4Rs if bound to G(i) inhibit adenylyl cyclase VI, whereas, if they are bound to G(s), they stimulate adenylyl cyclases VI and VII (Tucek et al., 2001). Alfonzo et al. (1998) show that membrane-bound guanylyl cyclase (GC) is regulated by muscarinic agents through two opposing signaling pathways. One involves the activation of GC through an M3R coupled to a pertussis toxin (PTX)–insensitive G protein. The other is mediated through a PTX-sensitive G-protein possibly coupled to an M2R. There can also be diVerences in the details of these cascades. In human neuroblastoma cells, the activation of ERKs 1 and 2 involves the sequential activation of PKC" ! Ras ! Raf ! ERK (Ras and Raf are types of GTPase) (Kim et al., 1999). The cascade described previously by Fukuhara et al. (2000) does not use Ras or Rho GTPases. In rat hippocampal CA1 cells, MAPK activity is modulated by both the PKA and PKC systems (Roberson et al., 1999).

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Furthermore, a wide variety of neuromodulatory neurotransmitter receptors (including MRs) use these two cascades. Also, PKC is a powerful modulator of phosphorylation of the transcription factor cAMP response element–binding protein (CREB). These workers found cross talk in the system in that MAPK activation is necessary for PKA coupling to CREB. CREB is implicated in synaptic plasticity and long-term memory. MR stimulation leads to a nonMAPK–dependent early (5m) phosphorylation of CREB and a late MAPKdependent induction of another transcription factor EGR-1 (Greenwood and Dragunow, 2002), which may also play an independent, or partially independent, role in memory processes. Platelet-derived growth factor decreases the number of M2Rs and the mRNA levels for M2Rs in HEL 299 cells. This is due to downregulation of M2R synthesis by means of an ERK pathway (Rousell et al., 1997). In pilocarpineinduced seizures, which are induced by excess M1R activity, Berkeley et al. (2002) have shown in mice that neuronal levels of ERK rise before the seizures start. SL327, an inhibitor of ERK activation, exacerbated pilocarpine-induced seizures. So the authors suggested that ERK might be neuroprotective in this situation. MAPK activity is also modulated by oxidative stress. In cultured striatal neurons, hydrogen peroxide (H2O2) powerfully increased levels of ERK 1/2 in a Ca2þ-dependent manner. H2O2 also strongly increases the release of arachidonic acid evoked by MR (as well as NMDAR) stimulation (Samanta et al., 1998). Cytokines, such as tumor necrosis factor alpha (TNF-) and interleukin-1, potentiate twofold to threefold the M5R-induced stimulation of arachidonic release by PLA2 (Wood et al., 2000). These cytokines also activate NF-B and MAPK pathways. Cytokines do not aVect other M5R-coupled pathways (e.g., PLC or intracellular Ca2þ mobilization). In many cases (e.g., growth factor and the -adrenergic receptor), receptor endocytosis is necessary to activate the MAPK/ERK pathway, however, not in all. Budd et al. (1999) showed that the ability of the Gq/11-coupled M3R to stimulate the ERK–1/2 pathway is not aVected by inhibition of its endocytosis. B. Other Kinases 1. Focal Adhesion Kinases (FAK) and Proline-Rich Kinases (PYK) These two tyrosine kinase families are involved in coupling neurotransmitter receptor activity to short-term and long-term changes in synaptic properties, cytoskeletal organization, and gene expression in neurons. Derkinderen et al. (1998) have shown in rat hippocampal slices that M1R and NR activity both stimulate PKC-mediated phosphorylation of FAKþ. Tyrosine phosphorylation of PYK2 (but not of FAKþ) was strongly enhanced by depolarization and

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hyperosmolarity. Thus, these two tyrosine kinases respond to diVerent activators. FAKþ is modulated by cholinergic input, whereas PYK2 responds to electrical and osmolarity parameters. Somewhat in contrast, Felsch et al. (1998) found that M1R activates PYK2 and that this is essential for the M1R-stimulated tyrosine phosphorylation of PYK2. This phosphorylation induces two cytosolic proteins, cSrc and Grb-2, to bind to PYK2. These workers also showed that the M2R controls the phosphorylation by PYK2 of the carboxy terminal of the Kþ channel Kvl-2. MR activation also promotes FAK phosphorylation during neuronal growth cone remodeling and neurite outgrowth (Linseman et al., 2000). This is mediated by the small GTPases Rho and Cdc42. Rosado et al. (2000a,b) note that oddnumbered MRs promote the tyrosine phosphorylation of a number of proteins. In a study of rat pancreatic acini cells, they showed that activating the M3R led to increased tyrosine phosphorylation of p125FAK, as well as of its substrates p130 (cas) and paxillin, by two pathways. One was dependent, and the other independent, of DAG-activated PKC activity and Ca2þþ. This eVect depended on the integrity of the cytoskeleton. Linseman et al. (1998) showed that the MR-activated tyrosine phosphorylation of FAK and paxillin is mediated by PI hydrolysis, and that this eVect also depends on the integrity of the actin cytoskeleton. The state of the actin cytoskeleton modulates muscarinic-evoked Ca2þ release. Depolymerization of actin filaments decreases, and polymerization increases this (Wang et al., 2002). Furthermore, in neuroblastoma cells, MR activation also stimulates tyrosine phosphorylation of FAK, Cas, and paxillin. This is mediated by a src-family kinase. However, in MR activation of ERK, src is not involved (Watcharasit et al., 2001). -Integrins play a crucial role as cytoskeletal anchorage proteins. In genetically engineered mice deficient in the 1 integrin gene, the normal muscarinic inhibition of L-type Ca2þ currents and activation of the Kþ current IK were both absent (Bloch et al., 2001). This was due to defective G-protein coupling. The authors conclude that cytoskeletal integrity is required for the correct localization and function of Gi-associated signaling domains. 2. ACKinase M3Rs activate Cdc42Hs-associated kinase-1 (ACK-1). The connecting chain does not involve Ca2þ or PKC but Cdc42, a Rho-family GTPase. This activity facilitates neurite outgrowth (Linseman et al., 2000). 3. Phospholipases Neurotransmitter receptors that couple to PTX-insensitive G (Gp) proteins typically evoke large increases in phospholipase C activity and subsequent release of Ca2þ from intercellular stores. In contrast, receptors that couple to PTX-sensitive G-proteins (Gi) usually generate weak PLC responses but strongly

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stimulate adenylyl cyclase. In transfected ovary cells, Carroll et al. (1995) showed that stimulation of M4Rs (coupled to PTX-sensitive G-proteins) strongly activates adenylyl cyclase but only weakly stimulates phospholipase C activity and the subsequent release of Ca2þ from intracellular stores. However, this latter eVect (together with muscarinic Ca2þ sensitive transcription of the c-fos gene) is greatly enhanced by the simultaneous activation the PTX-insensitive (PLC-activating) pathway by stimulation of adenosine receptors. Thus, the ability of M4R activation to induce PLC signaling strongly depends on the presence or absence of other PLC-activating agents. This mechanism may constitute a molecular coincidence detector that amplifies simultaneous PLC-activating signals involving cross talk between PTX-sensitive and PTX-insensitive G-protein pathways. 4. Phosphoinositol Signaling Activation of certain receptors leads to a postsynaptic signaling chain that autophosphorylates and so desensitizes the receptor. In transfected CHO cells, Budd et al. (2001) showed that activation of the M3R leads to two postsynaptic chains that cooperate synergistically to autophosphorylate the receptor. The first involves the activation of PKC by the diacylglycerol (DAG) arm of the PLC signaling pathway. This enzyme hydrolyses membrane lipids into two products— DAG and inositol 1,4,5 triphosphate (InsP3), both of which can act independently as further messengers. InsP3 releases Ca2þ from intracellular stores. The second independent route involves phosphorylation of the M3R on its third intracellular loop by casein kinase 1 (Budd et al., 2001). Johenning et al. (2002) have studied the distribution of various types of inositol 1,4,5-triphosphate receptors (InsP3Rs) and ryanodine receptors (RyRs) in diVerentiated PC12 cells. InsP3Rs type III are confined to the soma and function as gatekeepers for high-magnitude Ca2þ surges. This isoform is upregulated in neuronal apoptosis. Type I InsP3Rs are distributed throughout the cell and are related to signal initiation and repetitive signaling. RyRs types 2 and 3 in the soma serve as amplifying molecular switches, facilitating recruitment of the InsP3R type III–dependent pool. In the neurite, they decrease the probability of repetitive firing ( Johenning et al., 2002). The interaction of second messengers with their targets is constrained by microanatomical localization. In other words, a second messenger may fail to act, because it is unable to reach its receptor. An example is described by Delmas et al. (2002) In many excitable cells, M1Rs and B2 bradykinin receptors (both PLCcoupled) mobilize intracellular Ca2þ whereas in many neurons, only the B2R does this. In their experiments on sympathetic neurons, Delmas et al. (2002) found that whereas activation of both receptors rapidly produced DAG and InsP(3), only the InsP(3) formed by B2Rs has the ability to activate InsP(3) receptors, because it has microanatomical access, whereas the InsP(3) formed by M1Rs does not.

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V. Other Topics

A. Pathways from MRs to Gene Expression More detail on this matter has been provided by von der Kammer et al. (2001). They found 11 genes that were readily activated by MR stimulation within 1 hour. These were: —the transcription factors Egr-1/2/3, c-jun, Jun-D, and Gos-3 —the growth regulator hCyr61 —signaling factors NGFi-B and Etr101 —an unknown G-protein–coupled receptor-induced gene 2 (Gig-2) —the AChE gene These genes couple receptor responses to long-term neuronal responses. After 3 hours, a diVerent pattern emerged, including the expression of novel genes Gig-3 and Gig-4, as well as the LIM-only protein gene LM04. In neuroblastoma cells, MR activation induced expression of c-fos, fosB, c-jun, junB, and junD genes. The signal pathway included raised intracellular levels of Ca2þ. In the case of c-fos, fos-B, and junB, this was mediated by activation of PKC; in the case of c-jun and junB, it was mediated by Ca2þ/ calmodulin-dependent kinase II (Ding et al., 1998).

B. Cytoskeleton Stimulation of MRs by way of subunits activates the GTP-binding protein–coupled receptor kinase 2 (GRK2), which promotes the phosphorylation of tubulin, leading to the polymerization of microtubules (Haga et al., 1998). In addition to forming tubules, tubulin regulates certain G-protein–mediated signaling pathways. Both these functions depend on the ability of tubulin to bind GTP. Tubulin donates GTP to the G-protein G1 that regulates PLC (Popova et al., 2002). However, at high concentrations, tubulin will also inhibit PLC by binding (thus trapping) PLC’s substrate phosphatidylinositol 4,5-biphosphate (PIP2). Muscarinic agonists promote microtubule depolymerization and translocation of tubulin to the plasma membrane. Thus, tubulin, depending on local membrane concentration, may serve as a positive or negative regulator of phosphoinositide hydrolysis. The authors conclude that rapid changes in membrane phospholipid composition or in the cytoskeleton might modify neuronal signaling through such a mechanism. Activation of MRs leads to polymerization of microtubules, which, in turn, suppresses calcium channel current (Unno et al., 1999). In perisynaptic Schwann cells, ACh secreted by nerve terminals

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inhibits the neurotrophin glial fibrillary acid protein (GFAP) by means of MRs and Ca2þ. This results in cytoskeletal changes (Colomar and Robitaille, 2004). GFA levels in the pyriform cortex are also raised by AChE inhibitors (Zimmer et al., 1998).

C. More on Phospholipases 1. Phospholipase A In the glutamate synapse, PLA2 plays a key role in the post-NMDAR cascade [see my extensive review of this topic in Smythies (2002)]. PLA2 activity is important from a redox point of view, because it produces large amounts of potentially neurotoxic reactive oxygen species as a by-product. PLA2 also releases arachidonic acid (AA) from the hydrolysis of membrane lipids. This pathway may also play a similar role at the MR (Bayon et al., 1997; Lautens et al., 1998; May et al., 1999; Samochocki and Strosznajder, 1995; Shuttleworth, 1996; Shuttleworth and Thompson, 1998; Tence et al., 1994; Wood et al., 2000). In astrocytoma cells, this process requires PKC activity and G-proteins (Bayon et al., 1997). However, based on their studies of the distribution in murine brain of PLA2 mRNA and PLA2 immunoreactivity (which occur only in glia), Lautens et al. (1998) state that this pathway in the brain occurs only in astroglia. In a glioma cell line, DeGeorge et al. (1986) found that MR (but not NR) stimulation led to a rapid accumulation of AA derived from cyclooxygenase activity. In contrast, Tence et al. (1994) state that M1Rs activate PLA2 in murine striatal neurons in culture, leading to release of AA. They add that this reaction is positively regulated by PKC and Ca2þ. From experiments on brain synaptoneurosomes, Samochocki and Strosznajder (1995) concluded that MRs release AA in that tissue. This was almost completely inhibited by serotonin. In CHO cells, M3R activation stimulated PLC, AA release, and Ca2þ mobilization (May et al., 1999). They found that both PKA and PKC play a role in this process. Kjome et al. (1998) found that AA released by MR stimulation resulted in a homeostatic feedback inhibition of the MR. In contrast, Almeida et al. (1999) report that AA can have a positive feedback eVect on the MR in rat HC. In mammary adenocarcinoma lines, stimulation of MRs activated both cyclooxygenase (COX—a part of PGH synthase), leading to PGE2 synthesis and nitric oxide synthase (NOS) (Espanol et al., 2002). Both of these enzymes produce ROS as by-products. In ileal cells, the pathway for this same result was by means of PLC ! increased PI turnover and intracellular Ca2þ mobilization ! and activation of NOS and COX. Acetylcholine activates the HPA axis by

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means of COX-1–mediated PG synthesis (Bugajski et al., 2002a,b). Nitric oxide significantly inhibits this response (Bugajski et al., 1997). 2. Phospholipase C Phospholipase C is a phosphoinositide-specific enzyme. In the cerebral cortex and hippocampus, PLC-1 is involved in signal transduction by coupling by way of a G-protein to MRs (Kim et al., 1997). In contrast, in the cerebellum, PLC-4 acts through the metabotropic glutamate receptor (Kim et al., 1997). In transfected A9L cells, MR stimulation induces IP generation, AA release, and cAMP accumulation. The cAMP accumulation, however, was not due to a direct binding of the MR/G-protein complex with adenylyl cyclase or by way of the PLA2 and AA pathway, but is mediated by the PLC and PI pathway. This induced an increase in i.c. Ca2þ and calmodulin, leading to cAMP formation (Felder et al., 1989). In human upper airway ciliated cells, ciliary activation is produced by a muscarinic signal mediated by PLC (but not PLA) and its product prostaglandin (PG) E2 (Gayner and McCaVrey, 1998). But the carbachol-induced release of vasopressin in rat brain, after intraventricular infusion, is mediated by COX and PGD2. In smooth muscle cells, stimulation of M2Rs and M3Rs increases prostacyclin synthesis by means of COX, but the concomitant increase of cGMP synthesis is independent of COX ( Jaiswal et al., 1991). In cardiac cells, stimulation of M2Rs activates COX and increases prostaglandin synthesis ( Jaiswal et al., 1988). In colonic cells, stimulation of the M3R activates COX–2, which leads to increased PGE2 synthesis (Yang and Frucht, 2000). D. Adenosine Adenosine is an inhibitory transmitter in the CNS. Muscarinic and adenosine systems also interact. In rat cortical slices, stimulation of M3Rs does not aVect the basic rate of release of adenosine, but it does potentiate NMDA-evoked release of adenosine (Semba and White, 1997). In the unanesthetized rat brain, both nicotinic and muscarinic (as well as noradrenergic) stimulation increase extracellular adenosine levels (Bennett et al., 2003). In cultured rat hippocampal cells, M1R stimulation increases the AMPA-induced spontaneous excitatory postsynaptic currents (sEPSCs) (Bouron and Reuter, 1997). This is suppressed by a G-protein–dependent mechanism activated by presynaptic A1 adenosine receptors. In rat hippocampal CA1 neurons, stimulation of MRs produces a slow EPSP (Morton and Davies, 1997). This eVect is inhibited by adenosine acting presynaptically.

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E. Miscellaneous Items The AChE inhibitor pyridostigmine bromide (PB) lowers ATP levels in brain possibly because of its interference with mitochondrial function. This is associated with neuronal damage (Glass-Marmor et al., 1996). M4Rs (but not M1Rs or M1Rs), presumably after endocytosis, specifically bind to elongation factor, which is a GTP-binding protein essential for protein synthesis. M4R and elongation factor molecules colocalize in the soma and neuropil (McClatchy et al., 2002). The M4R molecule acts as a guanine-exchange factor for the elongation factor, suggesting a new role for the M4R in modulating protein synthesis. G-protein signaling is modulated by molecules called regulators of G-protein signaling (RGS). Stimuli such as activation of M2Rs that evoke neuroplasticity in neurons in the cortex, hippocampus, and striatum can rapidly induce mRNA for RSG2 (Ingi et al., 1998). This stimulates the GTPase activity of the -subunit of the G-proteins G(q) and G(i1). The authors conclude that this may provide a novel link between neuronal activity and signal transduction. The diurnal variation in activity of dopaminergic tuberoinfundibular neurons is modulated by MR stimulation by means of prostaglandin intermediaries (Chu et al., 2001). In rat olfactory bulb, M4Rs exert a bimodal control of cAMP synthesis through a sequence of events that may involve activation of Gi/Go proteins, synergistic interaction with Gs, and diVerential modulation of Ca2þ/CaMindependent and Ca2þ/CaM-dependent forms of adenylyl cyclase (Onali and Olianas, 1995). F. Reactive Oxygen Species Reactive oxygen species (ROS) play a prominent role in synaptic plasticity at the glutamate receptor (Smythies, 2002). There is also evidence that they play a role in muscarinic receptors. 1. Cholinergic Production of ROS In rat pheochromocytoma cells (PC12) stably expressing cloned M1Rs, activation of these receptors induces morphological changes and growth arrest. By use of this system, Mangelus et al. (2001) have shown that ROS act as second messengers in a number of MR-induced signaling pathways involving Ras, ERK, and p38, as well as activation of JNK, and an increase in the binding activity of the transcription factors NF-B and AP-1. Antioxidants in this system prevented the muscarinic-induced cellular changes. The authors indicate that the generation of ROS seems to be an early and critical event in these signaling cascades.

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MR stimulation, by raising i.c. Ca2þ levels, can also trigger PKC-mediated ROS production (in particular superoxide) in neuronal cells (Savolainen et al., 1998) and in neuroblastoma cells (Naarala et al., 1997). In smooth muscle cells, the pathway from MR stimulation to ROS production includes a PTX-sensitive G-protein, PI3-kinase, at least one tyrosine kinase, and opening the mitochondrial K(ATP) channel (Oldenburg et al., 2002). In pilocarpine-induced seizures, which are due to excessive muscarinic activity, the levels of the antioxidant enzyme superoxide dismutase (SOD) decreased in the HC, indicating the existence of oxidative stress (Bellissimo et al., 2001). The apoptosis induced by pyridostigmine bromide (PB) (a reversible inhibitor of cholinesterase producing raised levels of acetyl choline at its synapses) is mediated by ROS generated by activity at both MRs and NMDARs (Li et al., 2001). The source of ROS in the postsynaptic signaling chains initiated by MR activation may include COX and NOS (Smythies, 2002). 2. EVects of ROS on MR-Induced Events In human neuroblastoma cells, M3R activation induced increased phosphorylation in several adhesion-related proteins, including FAK, p130Cas, paxillin, and three shc adapter proteins. H2O2 inhibited this phosphorylation in all these proteins. In contrast, H2O2 increased the stimulation of the phosphorylation of the epidermal growth factor (EGF) receptor and Cbl induced by EGF ( Jope et al., 1999). Further experiments in human neuroblastoma cells by this group (Li et al., 1996) showed that ACh and H2O2 when given singly both induced increases in the transcription factors NF-B and AP-1. However, when they are given sequentially low doses of H2O2 had three eVects. They inhibited the increase in AP-1, increased PI hydrolysis produced by carbachol, and induced G-protein dysfunction. Activation of PI hydrolysis by the Ca2þ ionophore ionomycin was not aVected by H2O2, indicating that only certain forms of PI hydrolysis are aVected by H2O2. In heart cells, ROS aVect MR-induced eVects by reducing postsynaptic phosphoinositide (PI) turnover and increasing adenylyl cyclase inhibition. The result is an enhancement of the negative inotropic atrial response to ACh (Peters et al., 2001). In human neuroblastoma cells, the powerful oxidizing agent peroxynitrite (ONOO) also reduced MR-evoked PI hydrolysis, as well as decreasing the tyrosine phosphorylation of FAK and paxillin and increasing the tyrosine phosphorylation of p120Src (Li et al., 1998). ROS inhibit PI hydrolysis activated by metabotropic glutamate receptors (mGluRs), as well as by MRs in mice, but not that induced by activation of adrenergic receptors (De Sarno and Jope, 1998). 3. Redox Factors In the heart, GIRK potassium channels control cardiac contractility and excitability. Jeglitsch et al. (1999) have shown that these channels are redox sensitive. In cellular experiments, ROS, probably superoxide, markedly increased the

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current through these channels. In rat hippocampal CA1 neurons, carbachol induces a slow-onset long-term potentiation (LTPm) of reactivity to aVerent stimulation that is independent of the activation of NMDARs (Auerbach and Segal, 1997), as well as a non-NMDA–dependent tetanic LTP (NN-LTP). This eVect is abolished by low concentrations of H2O2. The authors suggest that the functional relevance of this is that the hippocampi of aged rats have higher endogenous H2O2 levels and lack LTPm and have markedly reduced NNLTP. These eVects in brain slices are reversed by catalase, which breaks down H2O2. Muscarinic receptors are diVerentially vulnerable to oxidative stress. In experiments on the ability of selectively transfected cells to clear the excess i.e. Ca2þ produced by ROS, M1, M2, and M4 receptors were more susceptible than were M3 and M5 receptors ( Joseph et al., 2002). Chronic hypoperfusion of the brain leads to hypoxia and oxidative stress. The resultant striatal neuronal cell damage is due mainly to a transient increase of NOS activity (which releases ROS as a by-product) and to a delayed-onset reduction of MR binding induced by low levels of glutathione (Tanaka et al., 2002). Low levels of another antioxidant—ascorbate—also reduce MR binding in salivary gland cell membranes (Sawiris and Enwonwu, 2000). In the bovine eye, the vasodilator responses induced by ACh and mediated by endotheliumderived hyperpolarizing factor (EDHF) are blocked by low doses of ascorbate (McNeish et al., 2002). Interestingly, in the human stomach, MR activation leads to a powerful secretion of ascorbate in gastric juice (Muto et al., 1997). Abu-Qare and Abou-Donia (2001) reported that a single dose of the anticholinesterase inhibitors (AChEis) sarin and PB led to increased urinary excretion of 3-nitrotyrosine and 8-hydroxy–20 -deoxyguanosine, which are markers of oxidative stress. Lipid peroxidation by ROS leads to a decrease of membrane fluidity and a kinetic MR malfunction (Ghosh et al., 1993). In rat experiments lasting 6 weeks, the inhibitory eVect of scopolamine on AChE can be reversed by increased levels of the antioxidant vitamins C and E in the diet (Lee et al., 2001a). In the brain in Alzheimer’s disease, an endogenous low molecular weight compound inhibits MRs by means of ROS-induced damage. This can be prevented by antioxidants (Fawcett et al., 2002). De Sarno et al. (2000) replaced the mitochondria in human neuroblastoma cells with platelet mitochondria from patients with Alzheimer’s disease. The chronic mild oxidative stress so induced in these cells led to two disturbances of MR function, namely an increase in PI signaling and a decrease in the binding of AP-1 to DNA. In mitochondria, ROS exert significant neurotoxicity by damaging the pathway for calcium extrusion leading to delayed calcium deregulation (DCD) and apoptosis (Nicholls et al., 1999). Mitochondria produce significant amounts of superoxide at complex III under conditions of high membrane potential ( m).

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G. Nitric Oxide In neuroblastoma and sympathetic neuron activation of MRs and NO, both rapidly raise cGMP levels; MRs do so by triggering Ca2þ mobilization, which stimulates NOS to produce NO, which stimulates soluble guanylate cyclase to produce cGMP. MR agonists also elicit a long-term (2h) suppression (LTS) of subsequent cGMP responses by means of activation of a nonspecific phosphodiesterase that lowers the concentrations of both cGMP and cAMP. LTS is induced by Ca2þ mobilization but is Ca2þ independent and is maintained by autophosphorylation of Ca2þ/calmodulin kinase II (CaMKII) (Trivedi and Kramer, 1998, 2002). MR activation stimulates NO release and cGMP formation in mouse neuroblastoma cells (Hu and el-Fakahany, 1993). The authors suggest that part of the NO generated stays within the same cell, and part may diVuse to neighboring cells. In CHO cells transfected with endothelial NOS (eNOS) and M receptor subtypes, stimulation of both M2Rs and M4Rs resulted in a marked activation of eNOS (Waid et al., 2000). This showed a transient peak (dependent on mobilization of iCa2þ stores and Ca2þ influx) and a sustained plateau that was dependent on Ca2þ influx only. Wang et al. (1997) confirm that M2R stimulation in transfected CHO cells activates NOS. In rat cortex, the ratio of catalytically active and inactive forms of NOS is driven by cholinergic input from the nucleus basalis of Meynert (HartlageRubsamen and Schliebs, 2001). The eVect of carbachol in producing NO is inhibited by arachidonic acid (AA). This treatment inhibits both the transient and the sustained increase in i.c. Ca2þ concentration evoked by muscarinic agonists (Linden and el-Fakahany, 2002). Thus, AA inhibits MR-activated NO production at the level of calcium mobilization. In transfected ovary cells, Parsons et al. (1999) report that exogenously applied NO regulates MR activity (PI formation), but endogenously produced NO does not. They suggest that this makes it unlikely that endogenously produced NO promotes negative feedback at the level of nNOS activation. In rat frontal cortex, M1Rs induce iNOS expression by means of the cascade ! PI ! PKC ! NOS, which results in the upregulation of M1R mRNA expression (Sterin-Borda et al., 2003). H. Tropins Several reported interactions between the muscarinic system and various tropins are important for synaptic plasticity. In the HC, NGF and BDNF are synthesized by neurons in an activitydependent manner (Knipper et al., 1994). Nerve growth factor (NGF ) plays an important role in maintaining inter alia the function of cholinergic neurons in the

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basal forebrain. In the reverse direction, cholinergic activity controls the NGF levels in rat cholinoceptive neurons in the cortex and HC. This is eVected by modulating the retrograde transport of NGF rather than by new synthesis (Rossner et al., 1997). In rat HC, Knipper et al. (1994) report that MR activation upregulates NGF and BDNF mRNA and NGF protein levels. Conversely, NGF and BDNF enhance ACh release from axon terminals. NGF also rapidly increases the high-aYnity choline transport into synaptosomes. Pesavento et al. (2000) present evidence that the eVect of NGF on long-term potentiation in the visual cortex is mediated by cholinergic receptors. In the HC, the levels of BDNF and NGF are largely controlled by activity in the glutamate and GABA systems. But, in experiments on rat HC, da Penha Berzaghi et al. (1993) showed that the muscarinic system also plays a role. Injections of the muscarinic agonist pilocarpine markedly increased both BDNF and NGF mRNA levels in the HC. French et al. (1999) report that activation of both MRs and NRs in the HC caused a transient increase in BDNF, NGF, and NT-3 mRNA levels but a marked rise only in the case of NR activation of NGF levels. Hohmann and Berger-Sweeney (1998) give a review of the complex morphogenetic interactions of cholinergic basal forebrain aVerents to the cerebral cortex and the neurotrophins BDNF, NGF, and NTB-4/5, possibly based on cholinergic interactions with diVerential gene expression. Cholinergic projections from the basal forebrain to the somatosensory cortex are necessary for BDNF inducement of rapid plasticity (Prakash et al., 2004). In the terminal fields in the basolateral amygdala (BLA) of axons from the nucleus basalis of Meynert in the rat, Moises et al. (1995) showed that NGF promotes the synthesis and release of ACh. This NGF-promoted presynaptic enhancement of the eVectiveness of cholinergic synapses was not accompanied by alterations in the properties of the resting membrane or in the intrinsic excitability of the BLA neurons. Pilocarpine-induced seizures (produced by excessive M1R activation) lead to increased levels of BDNF and neuropeptide Y expression (Poulsen et al., 2002). In contrast, in human neuroblastoma cells, stimulation of MRs inhibits signaling induced by EGF. This included PKC-mediated tyrosine phosphorylation of Cb1 and of the EGF receptor and complex formation between Shc proteins and Grb2 with the EGF receptor. In the reverse direction, activation of EGF receptors inhibited the MR-induced phosphorylation of FAK and paxillin. ROS, which are components of the EGF signaling cascade, mediated this cross talk. Thus, ACh and EGF have mutually inhibitory interactions (Zhang and Jope, 1999). Iwase et al. (2001) report that another tropin is hippocampal cholinergic neurostimulating peptide (HCNP), which enhances the synthesis of ACh in rat medial septal nucleus. The CA1 neurons in the HC contain high levels of the mRNA of the HCNP precursor protein. The expression of this mRNA is

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controlled by the reciprocal activity of cholinergic septal neurons and of glutamatergic hippocampal neurons. Furthermore, the activity-dependent and the constitutive expression of this mRNA may be controlled by diVerent routes involving Ca2þ influx by means of L-type Ca2þ channels and NMDARs, respectively (Iwase et al., 2001). I. Endocytosis Many receptors for neurotransmitters and neuromodulators, after stimulation, are subject to endocytosis into the interior of the cell and recycling back to the plasma membrane. This process plays a major role in the postsynaptic mechanism of the neuron and other cells. I have reviewed the roles of this process in the case of the glutamate receptor elsewhere (Smythies, 2002). Endocytosis can occur by three main mechanisms. The receptor may be engulfed in a clathrincoated pit (a process in which dynamin is involved) or in a calveolus or use neither. It is then taken into the interior of the cell. Here it is transferred to the endosome system from whence it may be traYcked to a proteosome for proteolysis or back to the plasma membrane for reuse. Endocytosis mediates several key functions, including desensitization and resensitization of receptors, control of receptor numbers, dephosphorylation of receptor proteins, repair of oxidative damage to membrane proteins and lipids, and redistribution of membrane between pruned and new synapses. 1. Endocytosis of the M1R In a series of experiments with CHO cells transfected with various M1R mutants, Shockley et al. (1997) showed that endocytosis and downregulation of the receptor are distinct events driven by diVerent cellular machinery. Stimulation of M1Rs induces the cross endocytosis of another receptor—the mGluR1 splice variants. This process requires PKC and Ca2þ calmodulin-dependent kinase (Mundell et al., 2002). 2. Endocytosis of the M2R In HEK-293 cells, M1–M4 receptors all internalize after stimulation, and all, except the M2R, use the clathrin or caveolae systems and rapidly recycle to the plasma membrane (Krudewig et al., 2000). The failure of the M2R receptor rapidly to recycle leads to a lengthy desensitization of M2R-coupled PLC, because the loss to the plasma membrane of the M2R is not compensated for by receptor recycling or receptor reserve. The M2R does recycle after stimulation but only very slowly (Roseberry and Hosey, 1999). This process has a component that requires new protein synthesis and a component that does not. This new protein does not consist merely of new receptor protein. Endocytosis of

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the M2R is unusual, because it does not use clathrin or calveoli (Hosey et al., 1999; Roseberry and Hosey, 2001; van Koppen, 2001; Vogler et al., 1999), yet dynamin may be involved in some way (Roseberry and Hosey, 2001; van Koppen, 2001; Werbonat et al., 2000) or may not (Hosey et al., 1999; Vogler et al., 1999). Delaney et al. (2002) found that the M2R is endocytosed by means of a pathway dependent on the ADP-ribosylation factor 6 (Arf-6)GTPase. This pathway quickly merges with the main clathrin-dependent pathway in early endosomes. Desensitization of M2Rs involves agonist-dependent phosphorylation of the receptor protein on the third intracellular loop by a GRK (Hosey et al., 1999; Tsuga et al., 1998a,b). In rat striatal interneurons, M2R stimulation leads to rapid endocytosis to the endosomes and particularly to the multivesicular bodies concerned with degradation (Bernard et al., 1998). Inhibition of AChE, either by drugs or genetic engineering, alters the traYcking of M2Rs. Acute inhibition by metrifonate causes increased endocytosis of these receptors to the endosome. In contrast, in AChE-deficient mice, impaired delivery of M2Rs from i.c. stores results (Decossas et al., 2003). 3. Endocytosis of the M3R In human neuroblastoma cells, M3R endocytosis and recycling play a larger role in receptor resensitization than they do in receptor desensitization (Edwardson and Szekeres, 1999). This resensitization may involve the dephosphorylation of the receptors in an endosome compartment. In this case, there is normally a receptor reserve of 50%. In human SH-SY5Y cells, M3R desensitization depends on two diVerent G-protein–coupled receptor kinases, GRK-3 and GRK-6, that use diVerent pathways (Willets et al., 2001). These authors suggest that GRK6 seems to act as a classical GRK uncoupling the receptor from the G-protein G(q/11). Conversely, GRK3 may cause desensitization independently from receptor phosphorylation by protein–protein interactions to inhibit PLC activity. The predominant MR in oligodendrocytes is M3R, which acts through the PI pathway. Stimulation of these leads to receptor desensitization of PI hydrolysis without endocytosis within 5 minutes. Later, endocytosis and downregulation of receptors occurs (Molina-Holgado et al., 2003). 4. Endocytosis of the M4R In rat striatum, stimulation of M4Rs leads to a rapid endocytosis of those receptors on the surface of the medium spiny neurons but has no eVect on the intracellular location of M4Rs in cholinergic interneurons (Bernard et al., 1999). NG108-15 cells express predominantly the M4R linked by means of a G-protein to adenyl cyclase. Stimulation of the M4Rs causes internalization and desensitization of the receptor. This eVect is mediated by G-protein–coupled receptor kinase GRK)-2 (Holroyd et al., 1999).

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During endocytosis, the M4R colocalizes with transferrin in the early endosome (Volpicelli et al., 2001), a process dependent on Rab5 activity. But each is directed to a diVerent part of the endosome—the M4R to the perimeter and interior of these vacuoles, transferrin to the perimeter only. A further study by this group (Volpicelli et al., 2002) showed that Rab 11a and myosin Vb are also involved in traYcking of the M4R. J. Arrestins According to Lee et al. (1998), alone among the MRs, the 2 variety depends on arrestin for the phosphorylation of its third intracellular loop required to initiate endocytosis This phosphorylation occurs at two ser/thr-rich sites on this loop—at P1 (286–290) and P2 (307–311). Phosphorylation at either site can support agonist-dependent endocytosis, whereas phosphorylation at site P2 is needed for the receptor to interact with arrestins (Lee et al., 2000a,b). Mundell and Benovic (2000) report that M1Rs and arrestins are connected in the sense that M1R stimulation induced endocytosis of arrestins to early endosomes. Arrestins also seemed to be involved in the desensitization of M1Rs. In a later article, the same group (Mundell et al., 2002) found that carbachol induced the translocation of arrestin from cytoplasm to plasma membrane in cells coexpressing mGluR1a or 1C but not in cells coexpressing mGlR1b. In experiments designed to detect peptide–peptide interactions, Wu et al. (1997) report that -arrestin binds tightly to the M3R protein. In cardiac cells, -arrestin plays a major role in the desensitization and other aspects of the function of the muscarinic Kþ channel (Shui et al., 2001). Gurevich et al. (1995) specify the structural elements in the arrestin molecule that determine its receptor binding specificity and selectivity. K. Miscellaneous Items In cardiac myocytes, MRs (type unspecified) are endocytosed by means of caveolae (Dessy et al., 2000; Feron et al., 1999) in which process NOS plays a role. MR endocytosis requires ongoing PI synthesis from phosphatidylinositol4,5-biphosphate mediated by PI4K (Sorensen et al., 1997, 1998). Astrocytoma cells have MRs coupled to PLC and -adrenergic receptors coupled to adenylate cyclase. Both of these reactions depend on PI synthesis for their endocytosis (Sorensen et al., 1999). Kinetic studies in human neuroblastoma cells showed that, in the case of the M3R, the same protein conformation is required for the generation of second messengers and for endocytosis. However, the evidence suggested that recycling is a pure constitutive process that does not depend on

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which agonist had triggered the endocytosis (Szekeres et al., 1998). Internalization of the MR is regulated by RhoA (Gampel et al., 1999). From experiments on a human dopamine cell line, Leng et al. (2001) report that stimulation of M1Rs and M3Rs leads to a transient release of membrane PLD2 from inhibition by -synuclein, which allows this lipase to participate in MR endocytosis. Chronic treatment of striatal neurons in vivo leads to increased endocytosis of MR2 and MR4 with a steep fall in their numbers in the plasma membrane and a rise in endosomes as well as at the sites of their synthesis (i.e., the endoplasmic reticulum nuclear membrane and Golgi apparatus) (Liste et al., 2002). L. Apoptosis A prominent eVect of neurotoxic AChE inhibitors is to induce apoptosis in neurons. Paradoxically ACh can often protect against apoptosis. 1. Neurotoxic EVects of MR Activation The toxic eVects of anticholinesterase inhibitors (AChEis) is mediated in part by damage to mitochondria, matrix swelling, and loss of ATP generation. Prominent among these agents are organophosphorus (OP) compounds such as sarin and pyridostigmine bromide (PB). Carlson and Ehrich (1999) measured the mitochondrial transmembrane potential psi(m). OPs induced an initial mitochondrial hyperpolarization followed by a depolarization. Pretreatment with carbachol significantly increased the hyperpolarization and delayed the depolarization. Pyridostigmine bromide (PB) is a reversible inhibitor of cholinesterase and leads to raised levels of Ach at its synapses. PB can induce apoptosis. This eVect is mediated by ROS generated by activity at both MRs and NMDARs (Li et al., 2001) and is associated with elevation of caspase activity and mitochondrial damage (release of cytochrome c). The authors conclude that MR-mediated ROS generation is an initiating factor in PB-induced apoptosis and that activation of the NMDAR is also directly linked to the response. The same group demonstrated that even a short exposure to PB could result in a prolonged (up to 30 days) apoptosis in rat cortical neurons in culture (Li et al., 2000). PB also induces a rapid reduction of mitochondrialbound hexokinase, an enzyme that is closely linked to mitochondrial oxidative metabolism (Glass-Marmor and Beitner, 1996). The intense overactivity of MRs during the seizures induced by pilocarpine leads to mitochondrial damage marked by a decline in function of complex I and complex IV components of the respiratory chain in rat CA1 and CA3 neurons (Kudin et al., 2002). This was associated with a lowering of the mitochondrial membrane potential in both hippocampal fields.

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The NMDA receptor antagonist phencyclidine (PCP) induces apoptosis in rat retrosplenial cortex (GriYths et al., 2000). This eVect is blocked by the muscarinic antagonist scopolamine, which suggests that ACh in this location promotes PCPinduced apoptosis. However, in the striatum, scopolamine does not have this eVect. PCP-induced apoptosis in the striatum is mediated by a corticosteroid system (GriYths et al., 2000). In HeLa cells that overexpress M1Rs, carbachol induces proteolytic cleavage of Rho-associated kinases (ROCK), a process involved in apoptosis (Ueda et al., 2001). Intermediary components in this eVect are G(11) and caspase. Thus, excess cholinergic activity could induce excess apoptosis by excess ROS production or by excess ROCK cleavage. 2. Neuroprotective EVects of MR Activation Activation of MRs protects against the apoptosis induced by various agents. In human neuroblastoma cells, DNA damage leads to activation of the proteolytic enzyme caspase-3, accumulation of p53, and apoptosis (De Sarno et al., 2003). MR activation prevented apoptosis, greatly attenuated caspase-3 activation but not p53 accumulation. H2O2 rapidly induces apoptosis by means of activation of caspase-3. This is prevented by pretreatment by muscarinic agonists. Rotenone is an inhibitor of mitochondrial complex 1 activity and induces apoptosis. This, too, was prevented by muscarinic stimulation. Curiously, muscarinic stimulation gave no protection to the apoptosis induced by staurosporine-activated apoptosis. Muscarinic protection against apoptosis produced by camptothecin was mediated by blockade of the release of mitochondrial cytochrome c associated with the bolstering of mitochondrial bcl-2 levels and blockade of the translocation of Bax to mitochondria. The latter eVect was also seen in the case of muscarinic protection against the apoptosis induced by H2O2 and rotenone. Thus, these results demonstrate that stimulation of MRs can oVer substantial protection against the induction of apoptosis by a variety of neurotoxic agents (De Sarno et al., 2003). PC12 cells deprived of support by tropic factors in serum undergo apoptosis. Activation of M1Rs inhibits this eVect by preventing activation of caspase-2 and caspase-3. In these cells, M1R stimulation also activates the MAPK/ERK and PI3-kinase signaling pathway, but these are not involved in the neuroprotective eVect of M1R stimulation against this type of apoptosis (Leloup et al., 2000). Lindenboim et al. (1995) also found that M1R activation protected against apoptosis in transfected PC12 cells deprived of growth factor. Sphingolipid metabolites, such as sphingosine and ceramide, play important roles in cell proliferation, diVerentiation, and apoptosis. In CHO cells, both these agents induce apoptosis (Lee et al., 2001b). They also inhibit the activation by muscarinic agents of PLD (but not PLC). Thus, it is possible that MR activation of PLD may protect against apoptosis in these cells.

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Cultured rat cerebellar granule cells undergo apoptosis when maintained in the presence of physiological concentrations of Kþ. This eVect is inhibited by activation of MRs, possibly type 3 (Yan et al., 1995). This eVect is not related to changes in glutamate release or i.c. Ca2þ levels. Calderon et al. (1998) have studied the mechanism of toxicity of purified AChE on neuronal and glial cells in vitro. They found that the apoptosis it produces was not related to its enzymatic action but to the fact that it promotes the assembly of -amyloid fibers into neurotoxic amyloid fibrils. The antioxidant vitamin Q10 has been reported to protect against apoptosis produced by excess levels of ROS in mitochondria (Alleva et al., 2001). In CEM-9F3 cells, vitamin Q10 protects against ROS-induced apoptosis by preventing oxidative damage and thus avoiding the activation of downstream eVectors such as neutral-sphingomyelinase, ceramide release, and caspase activation (Fernandez-Ayala et al., 2000). Tacrine is an anticholinesterase inhibitor that is used in the treatment of AD. Its beneficial eVects are thought to be due to raised levels of Ach in the brain (Sunaga et al., 1993). M. Neurotoxic Effects of Cholinesterase Inhibitors These agents like pyridostigmine bromide (PB) or sarin induce severe convulsion and death acutely. Even a single dose of induces long-lasting changes in the brain (Abdel-Rahman et al., 2002b), including mitochondrial damage and apoptosis. In rat experiments, 90 days after a single dose of sarin, the animals still showed a 29% inhibition of AChE activity in the cortex but a 20% increase in the brainstem. The cortex showed a significant decrease, and the brainstem a significant increase, in M2R receptor binding ( Jones et al., 2000). Henderson et al. (2002) followed a population of rats given sarin inhalations for 1, 5, and 10 days under normal and heat-stressed conditions. Observations were made at day 1 and at day 30. There were few changes at day 1 except for upregulation of brain mRNA of the cytokines IL-1, TNF-, and IL-6. After 30 days, there were no behavioral abnormalities or any changes observable by routine histopathologic findings in the brain. Also, there were few changes in the rats exposed to sarin alone. In those exposed to heat stress as well, however, after 30 days, M1R densities were decreased in the frontal cortex and HC; and M3Rs were upregulated after 1 and after 30 days in frontal cortex, striatum, and elsewhere. Sarin exposure by itself reduced AChE levels in the cortex and striatum. Sarin plus heat stress reduced AChE levels in the HC. Thus, the authors conclude that repeated exposures to very small doses of sarin can result in alterations in MR subtypes that may be associated with memory loss and cognitive dysfunction. Similarly, restraint stress and PB together (but not singly)

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induced disruption of the blood–brain barrier and widespread neuronal cell death in rats (Abdel-Rahman et al., 2002a). Repeated exposure to OP insecticides diVerentially downregulates mRNA expression of MRs in the brain. By in vivo studies in rats, Yagle and Costa (1996) showed that MIR mRNA was reduced in the HC with no change in the cortex, striatum, medulla, or cerebellum. M2R mRNAs levels were reduced in the hippocampus and medulla but not in the other three regions. M3R mRNA was reduced only in the cortex. Of these, the changes in M2 mRNA took the longest to recover. Soman causes excessive ACh activity, seizures, and a rapid increase in the levels of Fos, GFAP, and mRNA for AChE in the cortex. Zimmer et al. (1998) showed that this eVect is due to excess stimulation of MRs. It was accompanied by falls in the mRNA levels for ChAT and the vesicular ACh transporter (VAChT). A single dose of sarin also induces a longlasting expression of GFAP and vimentin genes (Damodaran et al., 2002). The authors suggest that this eVect may indicate damage to glia accompanied by compensatory proliferation and dediVerentiation. Much of the neurotoxicity induced by AChEis is mediated by means of GABA mechanisms. GrasshoV et al. (2003) showed that soman in the rat led to increased GABA release in the striatum mediated by excess cholinergic activity. In rat hippocampal slices, sarin inhibited GABA-mediated postsynaptic currents (PSCs) in CA1 pyramidal neurons but had no eVect on glutamate-evoked PSCs (Chebabo et al., 1999). However, this eVect of sarin was not related to its antiAChE eVect but to a direct interaction of the OP with MRs present on the presynaptic terminals of GABAergic neurons. Sarin had no eVect on the miniature postsynaptic currents mediated by either GABA or glutamate, indicating that sarin does not interact with GABA(A) or glutamate receptors. The authors conclude that the selective reduction by sarin of the action potential–dependent release of GABA in the HC can account for the seizures induced by this neurotoxin. This same group (Rocha et al., 1999) showed that the neurotoxin VX at low doses also reduced the amplitude of evoked GABAergic currents acting solely on the neurotransmitter release mechanism. NMDARs also play a major role in soman neurotoxicity. In animals that survive soman intoxication, lesions in the amygdala, piriform cortex, HC, and thalamus can be observed (de Groot et al., 2001). Antagonists of NMDARs were highly eVective in preventing the seizures and brain damage induced by soman. The authors conclude that cholinergic mechanisms are responsible for starting the seizures and that the seizures are maintained by the recruitment of other excitatory neurotransmitters and by the loss of inhibitory control that eventually leads to structural brain damage. Microperfusion experiments in vivo showed that soman leads to a major release of dopamine > GABA and aspartate in the rat striatum ( Jacobsson

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et al., 1997). These authors conclude that this major dopamine release may be as important as release of EAAs in the cascade of pathological events leading to the brain damage in the striatum seen after soman poisoning. N. Synaptic Plasticity ACh is involved in various aspects of synaptic plasticity. In the visual cortex, the M1R makes the predominant contribution among MR subtypes to the modifications of cortical circuitry that underlie plasticity (Gu, 2003). In transfected neuroblastoma cells in culture, stimulation of MRs induced increased neurite outgrowth and increased expression of the presynaptic protein synapsin-1 (De Jaco et al., 2002). This may be related to levels of early growth response gene-1. In the neuroblastoma cell line N1E-115, microinjection of ACh can induce the formation of filopodia and lamellipodia on growth cones. This eVect is mediated by cdc42Hs and Rac1, which are members of the Rho family of GTPases (Kozma et al., 1997). These authors conclude that neurite growth and retraction is controlled by competing pathways initiated by ACh using the pathway just described and lysophosphatidic acid (LPA), which operates by means of RhoA. The M3R modulates neurite outgrowth by activating Cdc42Hs-associated kinase (ACK-1) by means of a Fyn tyrosine kinase step (Linseman et al., 2001). Growth cones will turn in response to external DC electrical fields. This process was inhibited by MR and inhibited by NR activation (Erskine and McCaig, 1995). O. Gene Induction The muscarinic system plays a key role in the induction of many genes by several postsynaptic cascades. The MR-activated PKA-MAPK path plays a role in synaptic plasticity in the HC (Roberson et al., 1999). In the rat striatum, Wang and McGinty (1996b) report that MR activation in striatopallidal neurons inhibits the basal and D1R-stimulated dynorphin/substance P gene expression. It also facilitates the expression of the enkephalin gene in these neurons. In rat striatum, the muscarinic antagonist scopolamine did not by itself alter gene expression, whereas it profoundly augmented the expression of the two early genes c-fos and zif/268 induced by D1R activation (Wang and McGinty, 1996). Surprisingly, in the cortex, it attenuated both the basal and the D1Rstimulated expression of these early genes, indicating that ACh stimulates early gene expression in the cortex. The authors suggest that MR activity in the striatum may provide an activity-dependent inhibitory control of D1R activity.

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Activation of MRs leads to the promotion of the synthesis of AChE as a neuroprotective negative feedback mechanism. The pathway mediating this eVect is as follows. Stimulation of MRs activates PKC, which leads to the expression of the gene for the transcription factor Egr-1 and increased synthesis of this protein and its increased binding to and increased transcription from the AChE promotor region of DNA. This leads to increased AChE activity (Nitsch et al., 1998). In the adrenal medulla, activation of both MRs and NRs induces PNMT gene expression by means of the cAMP–PKA signaling mechanism (Wong et al., 2002). P. Cell Adhesion Molecules Cell adhesion molecules (CAMS), such as cadherin and integrin, play important roles in synaptic plasticity (Smythies, 2002). Some of this activity is modulated by the muscarinic system. In transfected CHO cells, cadherin-mediated cell adhesion is modulated by M3Rs and the small GTPase Rac1 (Ruiz-Velasco et al., 2002). M3R stimulation converts Rac1 to its GTP-bound form and alters the interaction between Rac1 and actin. In MR-transfected CHO cells, stimulation of M1Rs and M3Rs causes the cells’ endogenous cadherins to migrate to the plasma membrane and for the cells to adhere to each other (Shafer et al., 1999). This eVect is mediated by Ca2þ and PKC. In a carcinoma cell line, M3R activation also rapidly induces E-cadherin–mediated cellular adhesion mediated by PKC (Williams et al., 1993). Cells aVect chemical and electrical communication through gap junctions. Velasquez et al. (1997) reported that MR activation remarkably inhibited this process. Eosinophils adhere to airway cholinergic nerves and influence neuronal function by releasing granule proteins onto M2Rs. Coculture of eosinophils and IMR32 cholinergic neurons led to a significant increase in the release of eosinophil peroxidase and leukotriene C(4) (Kingham et al., 2002). IMR32 cells express the CAMs vascular cell adhesion molecule-1 and intercellular adhesion molecule-1 (ICAM-1). Eosinophil adhesion to IMR32 cells resulted in an ICAM1–mediated production of ROS by means of a neuronal NADPH oxidase and increased release of ACh from the IMR32 cells. Q. Hormones The muscarinic system is also involved in the physiology of various hormones. Cholinergic and dopaminergic systems in the preoptic anterior hypothalamus interact to regulate the release of luteinizing hormone during estrus (Cruz et al., 2001).

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In experiments on cell cultures of gonadotropin-releasing hormone (GnRH) hypothalamic neurons, Krsmanovic et al. (1998) showed that MR stimulation induced prominent peaks of GnRH release, each followed by a rapid decrease, a transient plateau, and a decline below basic levels. Nanomolar concentrations of ACh activated M2Rs and plasma-bound G(i) proteins that led to decreased cAMP production. Micromolar concentrations of ACh also caused an M1Rmediated increase in PI hydrolysis. These authors conclude that ACh modulates GnRH release through both M1R and M2R pathways by means of their respective PLC and adenylyl cyclase intermediates. In rat frontal cortex, activation of M1Rs stimulates the release of corticotropin-releasing hormone. The pathway was by way of a PTX-insensitive G-protein–activated adenylyl cyclase (Onali and Olianas, 1998). This same group identified the G-protein involved as G(q/11) and not G(i/o) (Olianas and Onali, 2000). In the mediobasal hypothalamus, IL–2 induces the release of corticotropin-releasing hormone by means of a sequential multisynaptic pathway involving both MRS and NRS—and NO (Karanth et al., 1999). In the pituitary, MR activation inhibits both (A) basal and (B) thyrotropininduced release of prolactin by two diVerent pathways (Pu et al., 1999). Both involved the PTX-sensitive cAMP/Ca2þ chain. But (A) was mediated by NO and (B) by i.c. calcium mobilization. Bayer et al. (1999) studied neurons in the diencephalon (zona incerta and perifornical regions of the hypothalamus) that produce melanin-concentrating hormone. They found that both MR and NR activation resulted in a rapid increase in the mRNA for this hormone. R. Vasopressin and Oxytocin Muscarinic mechanisms play a role in temperature, water balance, and blood pressure control. 1. Blood Pressure The rostral ventrolateral medulla (RVLM) has cholinergic mechanisms responsible for pressor responses (Kubo et al., 2000). Stimulation of the paraventricular nucleus of the hypothalamus causes an increase in arterial pressure by means of a cholinergic input from the lateral parabrachial nucleus to the RVLM (Kubo et al., 2000). Furthermore, the angiotensin system in the anterior hypothalamus is overactive in spontaneously hypertensive rats, which leads to raised levels of ACh release in the RVLM (Kubo et al., 2002). This chain leads to hypertension. There are two other cholinergic pressor systems. The most important consists of the central nucleus of the amygdala (AC). The other is composed of links

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between the lateral septum (LS), habenula, locus ceruleus (LC), and rostral VL medulla (Li and Ku, 2002). The AC widely connects with the other nuclei of the amygdala by means of its neurons that have corticotropin-releasing factor and substance P as neuromodulators. The second chain uses muscarinic receptors at each step. Li and Ku (2002) suggest that these two circuits are concerned in pressor responses induced by emotional stimuli. Experiments in rats involving local injections of muscarinic agents and measuring blood pressure responses and c-fos changes have identified brain areas concerned in this response (Li et al., 1997). The brain areas were the posterior, paraventricular, and supraoptic nuclei of the hypothalamus; the ventral premammillary nucleus; the central nucleus of the amygdala; the lateral septum; and the medial preoptic area. In spontaneously hypertensive rats, the hypothalamus exhibited increased levels of the mRNA for the M1R and decreased levels of the mRNA for the M4R (Wei et al., 1995). 2. Water Balance and Body Temperature The subfornical organ (SFO) is an important center for the regulation of vasopressin secretion and body fluid control. Xu et al. (2001a) showed by microinjection studies that cholinergic stimulation in the SFO induces water intake and c-fos expression in the anterior third ventricle region and in the vasopressin-containing neurons in the hypothalamus. This same group produced, by use of the same technique, more detail in this muscarinic system in the brain that controls water intake and vasopressin release. Besides the SFO, other nuclei lying along the lamina terminalis are involved. These include the organum vasculosum and the median preoptic nucleus. Further microdialysis studies showed that cholinergic stimulation in rats of the paraventricular nucleus (PVN) of the hypothalamus leads to a decrease in body temperature and an increase in water intake (Takahashi et al., 2001a,b). Similar stimulation in the supraoptic nucleus (SON) increased body temperature but had no eVect on water intake. This stimulation also led to increases in c-fos expression in the PVN, SON, LC, area postrema, and nucleus of the solitary tract. Microinjection of muscarinic agents into the preoptic area and anterior hypothalamus also leads to a fall in body temperature, increased water intake, and alertness (Mallick and Joseph, 1997; Takahashi et al., 1999c,d). In the rat medial preoptic area, only M2Rs may be involved in the modulation of thermoregulation and sleep (Imeri et al., 1996). In rat hypothalamus, melatonin inhibits the release of vasopressin and oxytocin by way of pathways using ACh, dopamine, and/or prostaglandins (Yasin and Forsling, 1998). In the SON, activation of MRs increases c-fos expression mainly in oxytocin neurons, whereas activation of NRs does so mainly in vasopressin neurons (Shen and Sun, 1995).

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S. T Cells T and B lymphocytes have M3Rs stimulation, which leads to Ca2þ signaling and c-fos expression (Fujii and Kawashima, 2000a,b, 2001). Human T-cell Jurkat line cells bear M1Rs and M2Rs. Stimulation of these modulate the production of the cytokine IL-2. Stimulation of the M1Rs influenced AP-1–mediated IL-2 promotion and induced the production of c-fos and c-jun protein. Stimulation of M1Rs also increased the activity of MAPK/ERK and JNK (Fujino et al., 2000). T. Learning Muscarinic systems are involved in learning-related activities, such as longterm potentiation (LTP) and long-term depression (LTD) and the encoding of information into long-term memory at a number of levels. The nucleus basalis of Meynert (NBM) evaluates sensory stimuli for their level of significance. ACh systems facilitate the processing of stimuli that are of behavioral significance at that moment (van der Zee and Luiten, 1999). The cholinergic nuclei supply the cortex, hippocampus, and amygdala with ‘‘strong facilitatory inputs at the critical moments when intensive information processing is required for short-term (e.g., hippocampus) or long-term (e.g., cortex) memory processes.’’ (van der Zee and Luiten, 1999). ACh provides the switch to promote information originating extrinsically over that arising intrinsically. It thus switches the hippocampus and neocortex from recall to learning modes (van der Zee and Luiten, 1999). In general, mRs modulate visual attribute processing, whereas NRs modulate arousal and selective attention to the visual task (Mentis et al., 2001). Elevation of ACh levels in the rat HC by the use of an AChEi enhances the induction of LTP by the inhibition of inhibitory GABAergic interneurons rather than by activating NMDARs on pyramidal neurons (Ye et al., 2001). Segal and Auerbach (1997) report two novel eVects of cholinergic stimulation on neuronal responses in the HC. The first was a fast-onset, short-lived increase in the cellular response to NMDAR activation. The second was a slow-onset, long-lasting increase in reactivity to aVerent stimulation of the muscarinic septohippocampal pathway that they called LTPm. The latter eVect was mediated by a postsynaptic M2 receptor and involves a rise in i.c. Ca2þ concentration and activation of both a tyrosine kinase and a ser/thr kinase. However, Parent and Baxter (2004) note a paradox in that, on the one hand, cholinergic activity in the HC is correlated with memory, and restoration of ACh in the hippocampus after disruption of the septohippocampal pathway is suYcient to rescue memory. However, on the other hand, selective ablation of cholinergic septohippocampal projections is largely without eVect on hippocampal-dependent learning and memory processes. The authors suggest that although hippocampal ACh is

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involved in memory in the intact brain, it is not necessary for many aspects of hippocampal memory function (Parent and Baxter, 2004). In monkeys, lesions of the cholinergic nucleus basalis of Meynert produce only modest or short-lasting impairments of visual discrimination learning whereas lesions of the medial septum/vertical limb of the diagonal band of Broca produce large and permanent defects of certain types of conditional learning (Harder et al., 1998). The authors conclude that rising cholinergic pathways sustain the functions of the target areas. In the case of the HC in humans, this is the analysis of information in a way that is pertinent to the formation of episodic memories. In the case of the neocortex, it is to process incoming information in a manner that is relevant to the cognitive processing of ongoing events and the acquisition of semantic knowledge. Pharmacological experiments on the rat hippocampal CA1 region led Kobayashi et al. (1997) to conclude that LTP there depends on the synergistic interaction between MRs and -adrenergic receptors. The MAPK/ERK pathway plays a critical role in learning and memory (Rosenblum et al., 2000). These workers showed that LTP in vivo requires ERKI/ II but not JNK in primary cortical cultures. This activation is Src-dependent, partly PI-3K- and Ca2þ-dependent, but PKC independent. M1 to M4 receptor subtypes in COS-7 cells can all induce ErkI/II activation using a similar signal transduction pathway. In the behaving rat, an unfamiliar taste activates ERKI/II in the taste area of the insular cortex culminating in the modulation of gene expression. This activity is mandatory for encoding the long-term (but not the short-term) memory of the taste (Berman et al., 2000). Multiple neurotransmitters are involved in this process. NMDARs, mGluRs, MRs, and -adrenergic and dopaminergic receptors all contribute to the acquisition of the memory of the new taste but not to its retrieval. Among these, only NMDA and MRs specifically mediate the activation of ERKI/II, whereas the -adrenergic function does not involve ERKI/II. DARs also regulate the basal level of ERKI/II activation (Berman et al., 2000). In vivo, the LTP potentiation in the insular cortex induced by stimulation of the basolateral amygdala requires NMDARs and is modulated by MRs ( Jones et al., 1999). The LTP also involves activation of ERKI/II. In the insular cortex, postsynaptic M1Rs and M3Rs (but not M2Rs) are involved in the acquisition of a conditioned taste response (Ramirez-Lugo et al., 2003). In rat perirhinal cortex, activation of M1Rs induced long-lasting depression (LPD) of synaptic transmission. This process is not dependent on NMDARS, synaptic stimulation, or PKC. It was, however, dependent on adequate i.c. Ca2þ stores and on protein synthesis (Massey et al., 2001). Mice lacking the gene for the M1R show a pronounced increase in locomotion and excess hyperactivity under stressful conditions but no cognitive impairment in a number of tests (Miyakawa et al., 2001). The authors suggest that the M1R may be less critical for learning

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than is generally supposed. However, it is possible that if a type of MR is developmentally absent from the brain, other MRs may take on part, or parts, of the function of the missing MR that they would not show in the normal brain. In the rat striatum, stimulation of M2Rs inhibits glutamate release from corticostriatal axon terminals and leads to a suppression of LTP. In mice, training in a spatial discrimination task induced increased numbers of MRs in the hippocampal CA1-2 pyramidal cell bodies and apical dendrites but a decrease in nonpyramidal neurons. However, mere exposure to the training board without learning was suYcient to lead to an increase in MR numbers in the CA3-4 neurons (van der Zee et al., 1995). In rat motor cortex, the induction of LTP, by layer I stimulation, in layer II/III horizontal connections requires activation of a diVuse cholinergic projection in the deeper layers (Hess and Donoghue, 1999). Hippocampal pyramidal cells (‘‘place cells’’) respond to the animal’s location in space. Muscarinic receptors mediate this function (Brazhnik et al., 2003). In the basolateral amygdala, MRs at cholinergic synapses regulate working memory but not short-term memory (Barros et al., 2002). Modulation of 48-hour retention memory functions in the BL amygdala requires both activation of M1 and M2Rs (Power et al., 2003). Spermidine can induce enhanced NMDA function by means of its modulatory site in the NMDAR channel. Kishi et al. (1998) showed that this could compensate for antagonist-induced dysfunction of rat hippocampal cholinergic- and mGluR-mediated neural function involved in working memory. Responses in the auditory cortex to a tonal conditioned stimulus show a specifically selective increase. This can be induced both by behavioral conditioning and by pairing a tonal stimulus with stimulation of the nucleus basalis. Miasnikov et al. (2001) showed that this involves the engagement of MRs in the auditory cortex. A recent study by Linster et al. (2003) discloses subtle modes of action of cholinergic systems in Freemanian-type learning reactions in the cortex. They suggest that ACh may regulate the dynamics of encoding and retrieval of information by regulating the magnitude of synaptic transmission at excitatory recurrent connections. Computer modeling suggests that the memory capacity of a realistic cortical network can be greatly enhanced if cholinergic modulation blocks transmission at synaptic connections of the association fibers during the learning process. The authors present experimental data from an olfactory cortex brain slice preparation showing that previously potentiated fibers show significantly greater suppression (presynaptic inhibition) by the cholinergic agonist carbachol than unpotentiated fibers. They conclude that a low level of suppression of nonpotentiated fibers during the learning process ensures the formation of self-organized representations in the relevant neural network, whereas a higher level of suppression of previously potentiated fibers minimizes interference

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between overlapping patterns. In a computational model of olfactory cortex, together these two phenomena reduce the overlap between patterns that are stored within the same neural network structure. The authors go on to suggest that these results further demonstrate the contribution of ACh to mechanisms of cortical plasticity and support a role for ACh in encoding new memories and enhancing response to salient sensory stimuli (Linster et al., 2003). U. Stress In a series of behavioral tests, File et al. (1998) showed that ACh induced a tonic anxiolytic response mediated by M1Rs, whereas the anxiogenic eVect of ACh is mediated by stimulation of presynaptic nicotinic autoreceptors to reduce ACh release. In a review of the literature, File et al. (2000) conclude that, in the dorsal HC, the postsynaptic 5-HT(1A) receptor mediates an anxiogenic response, whereas cholinergic tone (both muscarinic and nicotinic) mediates an anxiolytic response in the social interaction test. However, the authors found it surprising that nicotine also had an anxiogenic eVect in this test. The authors suggest that this is because of its further action in promoting serotonin release. They propose that the HC may control mainly the avoidance components of phobic anxiety with other regions, such as the dorsomedial hypothalamus, controlling the escape components. Blockade of the cholinergic system in the hippocampus by local injection of scopolamine augments the response of the HPA axis to restraint stress (Bhatnagar et al., 1997). Bugajski (1999) studied rats in a 21-day social crowding stress test. The result was that the increased release of ACh led to desensitization and downregulation of central MRs and to reduction of the HPA response to adrenergic agonists. Pare (2003) reports that stress hormones lead to the release of ACh and NE in the basolateral amygdala. V. Behavior Hippocampal activity that occurs in response to sensory stimulation is predominantly cholinergic. This can precede movement initiation and co-occurs with noncholinergic rhythm during ongoing behavior. In a series of ingenious experiments on the social behavior in two rats competing for one piece of food, Oddie et al. (1997) showed that muscarinic hippocampal is produced when the animal is showing behavior that involves sensory integration and social planning. Neocortical cholinergic aVerents are proposed to influence both selective attention and emotional processing. In a recent fMRI study of healthy adults, Bentley et al. (2003) used physostigmine to manipulate attention and emotionality to either faces with neutral or fearful expressions or house pictures. They found

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that physostigmine increased relative activity within the anterior fusiform gyrus for faces at attended, versus unattended, locations but decreased relative activity within the posterolateral occipital cortex for houses in attended, versus unattended, locations. Cholinergic enhancement augmented the neuronal response within the middle fusiform gyrus to fearful faces, whether at attended or unattended locations. In contrast, physostigmine influenced responses in the orbitofrontal, intraparietal, and cingulate cortices to fearful faces when faces occupied taskirrelevant locations. The authors suggest that these findings indicate that ACh may modulate both selective attention and emotional processes through independent, region-specific eVects within the extrastriate cortex. Furthermore, cholinergic inputs to the frontoparietal cortex may influence the allocation of attention to emotional information (Bentley et al., 2003). The circuit that mediates hypothalamic self-stimulation and ingestive behavior includes a muscarinic link in the VTA (Rada et al., 2000). W. Sleep Steriade (2004) has recently reviewed the role of ACh in certain aspects of sleep. His conclusions are that the two processes of activation in thalamocortical systems exerted by mesopontine cholinergic neurons are (1) a direct muscarinic depolarization associated with increased input resistance of thalamic relay neurons and (2) a disinhibition of the same neurons by means of hyperpolarization of inhibitory thalamic reticular neurons. Low-frequency ( limbic regions ¼ neocortex > thalamus (Abi-Dargham et al., 2000). 2. D2/3Rs Several groups have carried out PET scan studies of the distribution of D2Rs in primate and human brain. In humans, Okubo et al. (1999) found high levels in putamen and caudate nucleus, hypothalamus, red nucleus, and anterior and mediodorsal thalamic nuclei, as well as moderate levels in the globus pallidum, N.Ac., SN, subthalamic nucleus, amygdala, and other thalamic nuclei. The cerebral cortex had lower levels ranked temporal > anterior cingulate gyrus > parietal ¼ frontal > occipital. Another similar study in humans by Delforge et al. (2001) ranked D2R binding thalamus (2.9 pmol/ml) > temporal cortex (1.0 pmol/ml) > occipital cortex ¼ cerebellum (0.35 pmol/ml). The same group (Delforge et al., 1999) ranked B’ max levels in baboons thalamus (4.0) > cortex (0.32 – 1.90) > cerebellum (0.39). In macaque monkeys, Christian et al. (2000) found the following ranked distribution ratios: putamen, 29; caudate, 26; thalamus, 3.8; and frontal and temporal cortex, both at 1.7. In electron microscope studies, Hersch et al. (1995) found D2Rs in the dorsal striatum to be mainly presynaptic. In contrast, Flores et al. (1999) report in rat subthalamic nucleus, D1Rs, D2Rs, and D3Rs are postsynaptic, whereas D4Rs are wholly presynaptic. In a study of species diVerences (mouse, rat, guinea-pig, dog, rabbit), Levant (1998) found that the highest densities of D3Rs are found in the Islands of Calleja and in N.Ac., with moderate densities in the caudate nucleus in all species. The rabbit was unique in having high densities in the dorsal thalamus, posterior mammillary nucleus, and dorsomedial interpeduncular nucleus. The rat

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was unique in showing moderate densities in the molecular layer of cerebellar lobule X. Gurich and Joyce (1999) reported in the rat that the D3R is confined to mesolimbic structures and shows little overlap with D2R distribution. In contrast, in humans D2 binding sites were found in all areas studied by in situ hybridization receptor binding and mRNA level methods and showed much overlap with D2R distribution. They found the highest levels in humans in the striatum, N.Ac., external segment of the globus pallidus, SN, VTA, medial preoptic area, and tuberomammillary nucleus of the hypothalamus. D3Rs were abundant in the limbic striatum and its eVerent structures. Unlike the rat, in the human, D3Rs were virtually absent from the VTA. D3Rs were also found in the nucleus basalis of Meynert, dorsomedial and geniculate thalamic nuclei, mammillary nuclei, and the basolateral, basomedial, and cortical nuclei in the amygdala. The number of D2Rs in the brain decrease with age in females but not in males. This is not associated with diVerences in sex hormones (Kaasinen et al., 2000). 3. D4Rs The location of D4Rs in rat brain was studied by Wedzony et al. (2000) using immunocytochemical (Western blot analysis) methods. They found D4Rs in all regions of the prefrontal cortex (PFC) (cingulate, agranular/insular/orbital) and in all those cortical regions adjacent to the PFC (frontal, parietal, and piriform cortex). In subcortical areas, substantial levels of D4Rs were located in the striatum and N.Ac. In the cortex, the highest densities were in layers II/III and V/VI on both pyramidal output neurons and INs, especially on their dendrites. Using antireceptor antibodies Ariano et al. (1997) in rat brain found significant levels of D4R protein in frontal and parietal cortices with lesser amounts in the thalamus, globus pallidum, HC, and cerebellar vermis but only very low levels in the striatum (vide Wedzony et al., 2000). In rat N.Ac. shell, D4Rs are mainly extrasynaptic, with only a few at postsynaptic locations (Svingos et al., 2000). The location of the DA transporter (DAT) in human brain has been studied by Ciliax et al. (1999). They reported high levels, as one would expect, on most brain DA pathways along their entire somatodendritic and axonal domains, except for a small subpopulation of mesencephalic DA neurons and in all DA axons in the hypothalamus. The authors conclude that the latter groups must use some diVerent methods to control their extracellular (e.c.) DA levels. C. Basic Functional Anatomy The actions of DA on neurons are complex, indeed, depending on circumstances and are often biphasic. These will be covered in the next sections starting with basic issues and continuing with electrical responses and interactions with other transmitters.

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D1Rs link by way of Gs-protein and D2Rs by way of Gi-protein to adenylate cyclase. The former augments cAMP production, whereas the latter inhibits it (Lee et al., 2000). The D2R has two isoforms: L (long) and S (short). D2LRs are mainly postsynaptic, whereas D2SRs are mainly presynaptic (Borrelli, 2003). Virtually all striatal neurons carry both D1Rs and D2Rs, which can functionally interact (Aizman et al., 2000). Activation of D1Rs in both rat and human striatum converts associated D2Rs from a high to a low state of DA aYnity (Seeman and Tallerico, 2003). In the rat striatum, D1Rs synergize with D2Rs but not with D3Rs or D4Rs. This may be mediated by the interaction of receptors on separate striatal neurons in a manner not dependent on action potentials (LaHoste et al., 2000). The paraventricular nucleus (PVN) of the thalamus projects to both the PFC (deep layers) and to N.Ac. shell and plays a key role in the functional integration of limbic cortical and striatal circuitry. DA acts on the PVN–N.Ac. pathway by direct synaptic modulation but on the PVN–PFC pathway indirectly (Pinto et al., 2003). In cat motor cortex, DA inhibits the activity in both pyramidal tract and nonpyramidal tract motor neurons evoked by stimulation of the VL nucleus in the thalamus (Huda and Matsunami, 2003). In the PFC, there are two types of inhibitory interneurons—fast-spiking (FS) and non-fast spiking (non-FS). FS neurons synapse on the somata of pyramidal cells, whereas non-FS neurons synapse on their dendrites. DA depresses inhibitory transmission between FS neurons and pyramidal cells by way of D1Rs, but it enhances inhibitory transmission between non-FS neurons and pyramidal cells (not by way of D1Rs) (Gao et al., 2003). In the striatum, the glu input axons from the cortex and thalamus synapse on the spines of the output medium spiny neurons, whereas the DA input from the SN synapses mainly at the base of these spines (Tang et al., 2001). MRI studies in the rat show that activation of D2Rs increases activity in the VP and DM nuclei of the thalamus and in the hypothalamus. In PET studies in the baboon, Black et al. (2000) found that D1R agonists increased blood flow in the temporal lobe, including the amygdala, but reduced it in the thalamus, pallidum, and pons. Dopaminergic neurons possess D2-type autoreceptors that serve at least three functions: (1) somatodendritic autoreceptors decrease neuronal excitability; (2) terminal (i.e., presynaptic) autoreceptors reduce dopamine (DA) synthesis and packaging; and (3) terminal autoreceptors rapidly inhibit impulse-dependent DA release (Congar et al., 2002). DA neurons release DA not only from their terminal axons but also from their somata and dendrites (somatodendritic release). This requires the activity of synaptosome-associated protein of 25 kDa (SNAP–25) (Bergquist et al., 2002). There are many intrinsic factors that determine the strength of a synaptic event. These include the number of vesicles that open to discharge their contents, the geometrical relationship between these discharging vesicles and the target receptors, the nature of the contents of the synaptic space, and others. One of

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these is the number of neurotransmitter (NT) molecules released by a single vesicle during exocytosis. In DA neurons, these factors include (1) the rate of NT synthesis, (2) vesicle transport activity, (3) reuptake dynamics, (4) vesicle pH, and (5) fusion pore kinetics (i.e., what percent of a load is actually discharged). This factor is modulated by fusion pore shape, synaptic vesicle volume, and NT degranulation (Pothos, 2002). II. Electrical Reactions

A. Electrical Responses (Basics) DA can have both excitatory and inhibitory electrophysiological actions depending on circumstances. DA projections from the VTA to the PFC activate pyramidal neurons by way of D1Rs. PFC neurons have two resting states (1) a very negative resting membrane potential (‘‘down’’ state) interrupted by plateau depolarizations (‘‘up’’ state). In the rat, DA promotes the ‘‘up’’ state (Lewis and O’Donnell, 2000). Lavin and Grace (2001) confirmed this finding in rat PFC neurons and showed that the eVect is mediated by D1Rs. In the monkey PFC layer III pyramidal cells, D1Rs (but not D2Rs) have no eVect on the resting potential but increase excitability in response to depolarizing current steps injected at the soma (Henze et al., 2000). D1Rs (but not D2Rs) depolarize low-threshold spike INs in rat striatum suYciently to trigger action potentials (Centonze et al., 2002). In rat PFC layers II/III, DA inhibits IPSCs by activating presynaptic D1Rs (Gonzalez-Islas and Hablitz, 2001). In contrast, Gulledge and JaVe (2001) report that, in the rat medial PFC layer V, locally applied DA inhibits the activity of pyramidal neurons by two mechanisms (1) depression of spike generation and (2) depression of input resistance. The former is mediated by a GABAergic mechanism and the latter by a Naþ conductance. DA also triggers a longer-lasting increase in pyramidal cell excitability that is partly masked by the inhibitory action described previously. In the striatum, the action of DA on the medium spiny neurons depends on the state of polarization of their membranes. At resting potentials, DA promotes inwardly rectifying Kþ currents and so inhibits activity. On sustained depolarization of the spiny neuron, D1Rs now activate a Ca2þ current and so depolarize the cell (Reynolds and Wickens, 2002). B. Kþ-Related In the PFC, DA modulates the excitability of pyramidal cells both directly and indirectly by way of local circuit inhibitory GABAergic INs. In fast-spiking INs in rat PFC, DA induces a direct membrane depolarization, thus increasing

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their excitability. It does this by suppressing a voltage-independent ‘‘leak’’ Kþ current (by way of D1/D5Rs). Additional suppression of a slowly inactivating Kþ current leads to an increase in repetitive firing in response to depolarizing inputs. By this mechanism, whereby inhibitory INs are activated, DA can suppress persistent firing of pyramidal neurons (Gorelova et al., 2002). In rat SNpc neurons, DA inhibits their spontaneous firing by way of D2Rs, which activate a G-protein coupled to inwardly rectifying Kþ channels (Uchida et al., 2000). Isolated rat DA neurons in culture that have established self-innervating synapses (‘‘autopsies’’) release both DA and glu. In these cells, D2Rs autoreceptors at the presynaptic level inhibit glu-mediated synaptic transmission (EPSCs) by inhibiting inwardly rectifying Kþ channels and by more strongly inhibiting voltage-dependent Kþ channels. Both these Kþ channels are directly linked to the neurotransmitter release mechanism (Congar et al., 2002). In the rat SNpc, both DA and NE inhibit hyperpolarization-activated cationic I(h) currents and outward currents associated with an increase in I(Kir) conductance (Cathala and Paupardin-Tritsch, 1999). In contrast, in midbrain DA neurons, in two thirds of the cells, DA promotes I(h) conductance (Seutin et al., 2001). Dong and White (2003) describe three types of voltage-gated Kþ channels in rat medial PFC pyramidal cells—A type, I(D) type, and I(K) type. D1Rs inhibit only the I(D) type current by way of cAMP and PKA. In the transfected M8TG2 mesencephalon cell line D2S, D2L, D3Rs, and D4Rs couple to voltage-dependent Kþ ionic channels by way of diVerent G-proteins. Of these, the D2S, D2L, and D3 coupled systems increase the peak outward current, whereas the D4–G-protein coupled system decreases it (Liu et al., 1999). Glucose modulates DA neuronal activity and GABA axon terminal transmitter release in the rat SN by way of an ATP-sensitive Kþ channel. The direction of the modulation is both dose-dependent and time-dependent in complex ways (Levin, 2000). The author suggests that this may provide a mechanism whereby blood glucose levels modulate feeding behavior. C. Ca2þ Related DA can modulate the activity of calcium channels in various ways. There are six known types of Ca2þ channel, L-, N-, P-, Q-, R-, and T-, and one further type O-, has been proposed. Of these, L-acts mainly at postsynaptic sites and N-, P-, and Q- act at presynaptic sites supplying the Ca2þ ions needed for neurotransmitter release. Release can be triggered by Ca2þ ions from various sources. Stimuli not leading to the activation of voltage-operated Ca2þ channels can also elicit DA release, probably exocytotic in nature (Bonanno et al., 2000). Phillips and Stamford (2000) describe the complex pattern of involvement of N-, P-, and Q-type voltage-operated calcium channels (VOCCs) in diVerent parts

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of the rat striatum under conditions of both ‘‘regular’’ and ‘‘burst’’ firing. In cerebellar granule cells, D4Rs inhibit L-type Ca2þ channels (Mei, 1999). In human neuroendocrine cells, D3Rs inhibit P-/Q-type VOCCS by means of a percussive-sensitive G-protein and thus inhibit secretion by these cells (Kuzhikandathil and Oxford, 1999). In rat striatal enkephalin-expressing medium spiny neurons, D2Rs suppress L-type Ca2þ channels by way of the signaling cascade PLC ! IP3 ! calcineurin (Hernandez-Lopez et al., 2000). In rat subthalamic nucleus brain slices, D5Rs (but not D1Rs) promote activity in burst-firing neurons by way of the cascade G-protein ! PKA ! L-type Ca2þ channel. In this nucleus, presynaptic D2Rs reduce the impact of its two main inputs—from the cortex and from the globus pallidus—in this manner. Other postsynaptic DRs reduce intrinsic Kþ conductance that results in increased regular single spike firing (Baufreton et al., 2003). In medium spiny cells isolated from guinea-pig striatum, D2Rs produce only a very modest inhibition of Ca2þ conductance in normal cells but a very large inhibition in cells deprived of DA input by a 6-OH DA lesion in the SN. This eVect is mediated by a G-protein, PKC, and N-type VOCCS. Thus, this animal model of Parkinson’s disease involves the supersensitization of D2Rs (Stefani et al., 2002). In rat melanotrophic cells, DA can inhibit hormonal output. It does this by hyperpolarizing the cell membrane and by inhibiting high-voltage and lowvoltage–activated Ca2þ channels. On the other hand, the exocytotic machinery downstream from the Ca2þ influx is upregulated to compensate. Thus, the amount of hormone exocytosed per step depolarization can paradoxically not be diminished by DA, despite the inhibition of the Ca2þ current (Mansvelder et al., 2002). In rat neonatal (but not adult) midbrain, DA neurons exhibit spontaneous hyperpolarizations. These are mediated by increased Ca2þ release from i.c. stores, leading to increased free i.c. Ca2þ levels and the opening of apamin-sensitive Kþ channels (Seutin et al., 2000). D. Naþ Related In rat PFC layer V/VI pyramidal cells, older studies based on indirect evidence suggested that D1/D5Rs promote Naþ currents. However, more recent studies in rat striatum based on direct evidence show that D1/D5Rs inhibit rapidly inactivating Naþ currents (involving the Nav 1.1/1.2 Naþ channel) by way of cAMP and PKA. But these receptors do not aVect the persistent component of the Naþ current (involving the Nav 1.6/Naþ channel; Maurice et al., 2001). In rat HC, D1Rs reduce peak Naþ current by phosphorylating the -subunit of the Naþ channel by way of cAMP-dependent PKA. This is eVective primarily only when combined with depolarization or with activation of PKC.

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The phosphorylation of the -subunit of the Naþ channel is mediated at ser 573 on its intracellular loop L(I-II) by PKA anchored to A kinase anchoring protein–15 (AKAP–15). The reduction of peak Naþ current by PKC also requires phosphorylation of ser 554 of the -subunit and of ser 1506 in the inactivation gate. When ser 576 is phosphorylated by PKC, the increase in modulation by PKA requires phosphorylation of ser 687. This complex convergence of three distinct signaling pathways provides a mechanism for the integration of information from multiple signaling pathways (Cantrell et al., 1999; 2000). The D2 agonist -dihydroergocryptine activates presynaptic D2Rs by inhibiting voltage-gated Naþ channels. This agonist also modulates voltage-gated Naþ channels independently from D2Rs by some unknown mechanism (Neusch et al., 2000).

III. Interaction between Dopamine and other Transmitters

A. Dopamine ! Glutamate 1. Basics Sesack et al. (2003) review three anatomical sites of interactive modulation between the glu and DA: (1) dopamine regulation of pyramidal neurons in the prefrontal cortex by way of synaptic and extrasynaptic mechanisms and through indirect eVects mediated by GABA cells; (2) selective glutamate input from the prefrontal cortex or the brainstem tegmentum to diVerent populations of dopamine cells in the VTA; and (3) within target regions (the basal ganglia, prefrontal cortex, and amygdala) by way of convergent synaptic or extrasynaptic regulation of common neurons. 2. Presynaptic DA can both increase and decrease glu release. In rat and mouse olfactory bulb, D2Rs presynaptically inhibit glu olfactory nerve terminals (Ennis et al., 2001), and in the striatum, DA also inhibits glu release (Tang et al., 2001). In ferret PFC, DA inhibits the microcircuitry composed of excitatory glu transmission between pyramidal cells by a presynaptic mechanism but has no eVect on excitatory glu transmission directed at GABAergic INs (Gao and GoldmanRakic, 2003). In contrast, in rat HC, D1Rs presynaptically promote glu release by way of PKA (Bouron and Reuter, 1999), and in rat prelimbic cortex layer V/VI pyramidal cells, D1Rs presynaptically increase the frequency of spontaneous EPSCs by way of PKA and PKC (Wang et al., 2002).

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3. Postsynaptic Excitation (D1Rs) In cultured striatal neurons, D1Rs potentiate AMAP-induced EPSCs by way of PKA-mediated phosphorylation of the AMPAR (Price et al., 1999). In rat and mouse striatum, D1Rs potentiate NMDA-evoked responses. This potentiation is diminished by simultaneous activation of D2Rs and by activation of protein phosphatase 1 (PP–1). These reactions are eVected by way of the ubiquitous DARPP–32 pathway. In the same cell, D1Rs can potentiate NMDAevoked currents and inhibit GABA-induced currents (Flores-Hernandez et al., 2002). In rat PFC layer V pyramidal cells, D1Rs potentiate NMDAmediated excitability by way of a PKA-dependent pathway and Ca2þ-dependent mechanisms (Wang and O’Donnell, 2001). In rat PFC, Gonzalez-Islas and Hablitz (2003) confirm that DA potentiates EPSCs (produced by both AMPA and NMDA) by way of a cascade the involves Ca2þ ! activation of PKA ! Ca2þ/calmodulin-kinase II (CaMKII). In neonatal rat N.Ac., D1Rs increase the phosphorylation of the AMPAR GluR1 subunit at the PKA phosphorylation site ser 845. This eVect is inhibited by DR2s. DA had no eVect on phosphorylation at ser 831 of the AMPA subunit. This is the site of action of PKC and CaMKII. Phosphorylation here increases AMPA channel conductance. A similar result was reported by Snyder et al. (2000) in mouse neostriatal slices. D1/D5Rs in rat ventral HC exert a strong excitatory tone on NMDAR-mediated motor activation (general motor activity, locomotion, and rearing behavior) (Gimenez-Llort et al., 2002). 4. Postsynaptic Inhibition In rat striatal slices, D1Rs and D2Rs (and NMDARs) all inhibit CaMKII activity. DA, however, markedly increases PKA activity (Tang et al., 2000). In rodent motor cortex, DA inhibits and glu potentiates the spontaneous firing rate of pyramidal tract neurons. Co-application of DA inhibited the increase induced by glu (Awenowicz and Porter, 2002). In CA1 pyramidal neurons, D2Rs inhibit NMDA excitatory transmission by increasing Ca2þ inflow, leading to Ca2þ-dependent inactivation of NMDA receptors. This eVect depends on the ‘‘transactivation’’ of platelet-derived growth factor- (PDSFR ) (Kotecha et al., 2002). DA lowers the threshold for the induction of long-term depression (LTD) in rat PFC glu transmission by a mechanism involving cooperative postsynaptic depolarization between groups I and II mGluRs and DRs mediated by way of MAP-K (Otani et al., 1999). D1Rs also inhibit glu (NMDA)-induced inward currents in rat dorsal striatum medium spiny neurons (Lin et al., 2003). In cat motor cortex, iontophoretically applied DA inhibited (by way of both D1Rs and D2Rs) pyramidal tract neuronal responses to activation of both thalamocortical and callosal inputs (Huda et al., 2001).

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5. Miscellaneous EVects In rat caudate/putamen patch-like regions, D2Rs related to aVerents from the SN are strategically placed at both presynaptic and postsynaptic sites close to glu axons from the PFC to allow close modulation of their activity (Wang and Pickel, 2002). Mutant mice completely lacking D4Rs show cortical hyperexcitable brain glu systems expressed as an increase in both spontaneous synaptic activity and paroxysmal discharges (Rubinstein et al., 2001). In rodent PFC pyramidal cells in culture, D4Rs inhibit NMDAR-evoked currents. This process involves the chain PKA (inhibition) ! PP1(activation) ! CAMKII (inhibition) (Wang et al., 2003). Complex interactions between glu (NMDA), GABA, and DA systems in cultured medium spiny neurons from rat striatum are described by SchoVelmeer et al. (2000). NMDAR activation causes a slow action potential–independent eZux of GABA. The resulting entry of Naþ through the NMDA channel reverses the direction of action of the Naþ-dependent GABA transporter. Activation of G-proteins linked to D1Rs also induces GABA release by way of its transporter but by a diVerent mechanism. This is the cAMP-dependent inhibition of Naþ/Kþ-ATPase that results in the accumulation of i.c. Naþ, reversal of the GABA transporter, and potentiation of NMDA-induced GABA release. In transfected CHO cells, the DAT mediates a DA-dependent apoptotic signal that is independent of DA uptake into the cell (Liu et al., 2001). DA protects against glu neurotoxicity in a number of ways. Dopamine is a potent scavenger of reactive oxygen species (Liu and Mori, 1993). D2Rs can protect against glu-induced apoptosis (Bozzi et al., 2000). D2Rs activate the synthesis of i.c. antioxidant enzymes (Sawada et al., 1998). DA may be endocytosed together with its D1 receptor and play a role in the intracellular O’Brien cycle in which cycling between dopamine and dopamine quinones and between ferric and ferrous iron converts superoxide into less toxic products (H2O2 and water) (see Smythies [1999a, b] for details). D2R agonists protect neurons against excessive Ca2þ influx, NO, and ONOO-induced components of glu toxicity by activating phosphoinositide 3-kinase (PI-3K). The prosurvival Akt-protein (an eVector of PI-3K) and the antiapoptotic protein Bcl-2 are also involved in the neuroprotective eVect of DA (Kihara et al., 2002). DA is also easily oxidized to highly neurotoxic o-quinones. This reaction may be involved in the pathogenesis of Parkinson’s disease (Smythies, 1996, 1999a, b). B. Glutamate ! Dopamine 1. Presynaptic Group 3, and to a lesser extent group 2, mGlu RS presynaptically regulate DA release and uptake in rat N.Ac. in complex biphasic dose-dependent and mutually interactive ways (Hu et al., 1999). In rat SN, group 2 and 3 mGLURs

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also presynaptically inhibit DA release onto DA neurons (Katayama et al., 2003). In guinea-pig striatum, activation of AMPARs inhibits DA release by a presynaptic mechanism despite the fact that DA terminals here lack ionotropic glu receptors. The mechanism involved is through AMPA-evoked postsynaptic H2O2 release. This acts as a retrograde messenger to inhibit DA release by activation of sulfonylurea-sensitive Kþ channels. This eVect is opposed by simultaneous activation of GABA(A)Rs (Avshalumov et al., 2003). In the HC of freely moving rats, NMDARs inhibit DA release. The NO donor SNAP has a biphasic eVect on this response—a low dose inhibits and a high dose potentiates NMDAR-initiated DA release (Segieth et al., 2000). In contrast, in freely moving rats, stimulation of the direct glu projection from the basolateral amygdala to N.Ac. increased DA eZux by 25%. Stimulation of the central nucleus of the amygdala had no such eVect (Howland et al., 2002). In freely moving rats, NMDARs inhibit DA release in the frontal cortex and dorsal raphe. This process is modulated by NO, apparently by diVerent mechanisms in each brain locus (Smith and Whitton, 2001). 2. Postsynaptic In cultured striatal neurons, group 1 mGluRs (but not group 2 or 3) potentiate the cAMP production induced by D1Rs by way of PKC (Paolillo et al., 1998). In rat midbrain slices, mGluRS induce IPSPs in DA neurons. This initiates the signaling chain—release of InsP(3) ! release of Ca2þ from i.c. stores ! activation of small conductance Ca2þactivated Kþ channels (Morikawa et al., 2000). In rat SN, group 1 mGluRs also inhibit DA neurotransmission postsynaptically by opening Ca2þ-sensitive Kþ channels (Katayama et al., 2003). The DA and glu systems also interact at the level of traYcking of receptors from the cell interior to the external membrane. In rat striatum, NMDARs recruit D1RS from the cell interior to the plasma membrane (Scott et al., 2002). Likewise, in primary cultures of rat N.Ac. neurons, D1Rs increase the surface expression of the GluR1 subunit of the AMPAR (Chao et al., 2002). NMDA receptors also regulate D1 receptor function by means of a direct protein–protein interaction mediated by the carboxyl tail regions of both receptors. This allows NMDA receptors to increase plasma membrane insertion of D1 receptors and provides a novel mechanism by which the activation of NMDARs upregulates D1 receptor function (Pei et al., 2004). 3. Miscellaneous In rats, glutamatergic eVerents from the ventral subiculum project to N.Ac., where they excite GABAergic neurons. The latter project to the ventral pallidum, where they inhibit a second GABA projection—to the DA neurons of the VTA. Thus, glu eVerents from the ventral striatum stimulate activity in DA neurons in the VTA by this disinhibitory polysynaptic pathway. This results in increased DA

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release in N.Ac. and elsewhere. The ventral subiculum also maintains a steady-state basal level of activation of VTA DA neurons (Floresco et al., 2001). DA neurons in the SNpc and in the VTA release DA from their somata and dendrites. In the guinea-pig SNpc, this release is inhibited by glu input acting by means of inhibitory GABA INs. In contrast, in the VTA, the timing and/or strength of the synaptic input determines whether the net eVect on DA release is excitatory or inhibitory (Chen and Rice, 2002). The cystine-glutamate antiporter exchanges extracellular cysteine for intracellular glutamate. This releases glu into the extracellular space. It also plays a key role in the antioxidant defenses of the cell, because cystine is the metabolic precursor of the principal intracellular antioxidant glutathione. This transporter is inhibited by extracellular DA (which eVect is itself inhibited by group 2 and 3 mGluRs) and by group 2 mGluRs by means of a cAMP-dependent protein kinase mechanism (Baker et al., 2002). Chuhma et al. (2004) describe a novel interaction between dopamine and glutamate that depends on the fact that many dopamine axons secrete dopamine from preterminal sites but secrete glutamate from their axon terminals. These authors state that dopamine neurons are thought to convey a fast, incentive salience signal, faster than can be mediated by dopamine. They propose a resolution of this paradox, which is that midbrain dopamine neurons exert fast excitatory actions by means of their terminal glutamatergic synaptic connections to accumbens neurons. They conclude that this dopamine neuron glutamatergic transmission may be the initial component of the incentive salience signal. C. Dopamine and GABA 1. Anatomy Gonzalez-Hernandez and Rodriguez (2000) give a detailed account of the chemical anatomy of DA and GABA cells in the rat SN, including their content of CCK, calretinin, calbindin, parvalbumin, and NOS. They recognize five types of DA cell and five types of GABA cell that have various distributions in the various subdivisions of the SN. In both rat and monkey cortex, DA terminals synapse directly on GABA dendrites (Sesack et al., 1995). In rat striatal and N.Ac. cultures, D1Rs and D2Rs coexist on presynaptic varicosities and on cell bodies but not on axons (Wong et al., 1999). 2. Presynaptic In rats, DA inhibits GABA(B) transmission onto DA neurons by an ‘‘unconventional’’ presynaptic mechanism not involving D1Rs or D2Rs (Federici et al., 2002). In rat striatum, inhibitory D2Rs on the terminals of GABAergic aVerents to cholinergic INs downregulate GABA release by blocking N-type

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Ca2þ channels by means of NEM-sensitive G-proteins (Momiyama and Koga, 2001). In the striatum of freely moving rats, an increase in endogenous GABA levels facilitates Kþ-stimulated, but not basal, DA release (Galindo et al., 1999). 3. Postsynaptic In rat PFC pyramidal cells, D4Rs downregulate GABA(R) channel activity by inhibition of PKA and the subsequent stimulation of PP1. For this reaction to occur, PKA and PP1 must form a complex on the scaVolding protein Yotaio that targets them to the vicinity of GABA(A)Rs (Wang et al., 2002). 4. Functional Anatomy In rat PFC pyramidal cells, DA has a bimodal eVect on GABAergic IPSCs. D2Rs induce an initial abrupt reduction, whereas D1Rs induce a late slow increase of these IPSCs. D1RS act by increasing the intrinsic excitability of INs and their axons, whereas D2Rs act by inhibiting presynaptic GABA release and the postsynaptic response to GABA (Seamans et al., 2001). In this way, these authors suggest that D2R-related activity, which reduces inhibition, allows multiple representations, both weak and strong, to be activated closely linked in time. This is followed by an upsurge in D1R-related activity that increases lateral inhibition and so serves to focus activity on task-related items. Thus, activation of DARs may first promote an exploration of the input space (following the receipt of an unfamiliar or unexpected stimulus [i.e., ‘‘Something important has occurred—what and where is it?’’]). This is followed by a transition to a state in which focus turns to a limited subset(s) of representations that take over complete control of PFC output, allowing the weaker representations to die out (i.e. ‘‘So that is what is important—so now to deal with it.’’). D. Dopamine and Acetylcholine 1. DA—ACh In rat striatal synaptosomes, D2Rs modulate the Ca2þ-dependent component, but not other components, of spontaneous ACh release (Sanz et al., 2000). In rat corticostriatal slices, DA increases the excitability of cholinergic INs by a D1R-related polysynaptic mechanism. This is accompanied by a D2R-mediated presynaptic inhibition of GABAergic and cholinergic IPSCs (Pisani et al., 2000). In mouse corticostriatal slices, stimulation of corticostriatal fibers leads to an early glu-mediated depolarization of cholinergic neurons followed by later polysynaptic GABA-mediated IPSPs. Activation of D5Rs, but not D2Rs, is necessary for the Ca2þ-dependent expression of LTP in these neurons (Suzuki et al., 2001). The glu excitatory input to the VTA synapses on both DA and GABA cells. These two types of synapse show marked diVerences. In response to repetitive stimulation,

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the former exhibit LTP, whereas the latter exhibit LTD. When exposed to NMDA, the former develop LTP, whereas the latter do not (Bonci and Malenka, 1999). 2. DA—>5-HT In rat dorsal raphe, DA depolarizes 5-HT neurons by activating a nonselective cation current and not by inhibiting a Kþ current (Haj-Dahmane, 2001). 3. Other Interactions In rat SNpc, adenosine 1Rs (A1Rs) and D1Rs interact. The inhibition exerted by A1Rs on striatonigral transmission is due to blockade of the facilitation resulting from activation of D1Rs (Floran et al., 2002). In mouse transfected fibroblasts, adenosine by means of A1Rs inhibits D1R-mediated signaling by forming functionally interacting heteromeric complexes (Gines et al., 2000). In freely moving rats, D2Rs (but not D1Rss) promote the release of the endogenous cannabinoid anandamide. This constitutes an inhibitory feedback mechanism countering DA-induced facilitation of motor activity (GiuVrida et al., 1999). E. Interactions between Dopamine and Nitric Oxide The ubiquitous signaling molecule nitric oxide (NO) plays several roles in the dopamine system. A recent extensive review of this topic (West et al., 2002) concludes that NO generally facilitates both tonic and phasic DA neurotransmission in the striatum. In rat striatum, NO acts biphasically on DA levels. High levels of NO produce high levels of the neurotoxin peroxynitrate (ONOO-), which lowers DA levels by oxidation. Lower concentrations of NO increase the levels of DA (as well as 5-HT) probably by a cGMP-dependent mechanism (Trabace and Kendrick, 2000). In rat ventral HC, endogenous NO lowers extracellular levels of DA and 5-HT, in this case probably not by cGMP (Wegener et al., 2000). In rat HC slices, D1/D5Rs induce a sustained enhancement of glu EPSCs mediated by NO acting at a presynaptic level (Yang, 1999). In rat striatum, NO tone, mediated by NOS and INs activities, regulates the striatonigral projection neurons and thus modulates (increases) the response of DA neurons to input from the cortex by way of the striatum (West and Grace, 2000 q.v. for further complexities in this arrangement). In rat N.Ac. (both core and shell), DA and NOSþ terminals converge onto a single spiny dendrite and spine. In this system, NO also potentiates DA release (Hidaka and Totteredell, 2001). In rat striatum, endogenously produced NO increases DA release by action on the DAT (Kiss et al., 1999).

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NO also exerts a biphasic modulation of DA neurotoxicity. The toxicity of MPPþ (the active metabolite of MPTP that induces a model Parkinsonian syndrome) is mediated by the hydroxyl radical (OH). NO formation is necessary for this eVect. However, NO is not involved in MPPþ-evoked DA release (Rose et al., 1999). Lipopolysaccharides induce cell death by activating NOS (Iravani et al., 2002). Furthermore, NO oxidizes DA to neurotoxic dopamine quinones (Antunes et al., 2002) that link covalently to proteins and produce mitochondrial damage. DA quinones are mandatory intermediates in the synthesis of neuromelanin and probably play a key role in the pathology of Parkinson’s disease (Smythies, 2004). DA can also be metabolized to 6-nitrosodopamine, which is also a neurotoxin (Nappi and Vass, 2001). NO can also play a neuroprotective role and protects the DA system from iron-induced oxidative stress in rat brain (Lin, 1999).

IV. Dopamine-Induced Signaling Cascades

A. G-Proteins DiVerent DRs engage with diVerent postsynaptic signaling cascades. A single DR can also connect with a number of such cascades. In transfected Chinese hamster ovary (CHO) cells, D2Rs can interact with diVerent G-proteins to induce a number of responses: (a) inhibition of adenyl cyclase, (b) modulation of Kþ currents, and (c) potentiation of Ca2þ-evoked arachidonic acid (AA) release. Activation of PKC diverts D2R signaling from pathway (a) to pathway (c) (Di Marzo et al., 1993). In rat anterior pituitary cell membranes, D2Rs couple by means of G-proteins Gi1/2 to PLC and inhibit this enzyme (Rasolonjanahary et al., 2002). In rat midbrain DA cells, DA, by means of Gi/o proteins, inhibits DA cell firing, whereas MR1 and NMDARs by means of Gq/ll proteins increase the firing of these cells (Gronier and Rasmussen, 1999). In GH4ZR7 lactomorph cells in rat pituitary, D2SRs inhibit cAMP production and so block the phosphorylation of MAPKs by means of Gi/o proteins. But when coupled to G(I)3/G (o) proteins in this system, D2SRs inhibit MAPK by a diVerent pathway (i.e., by c-Raf and -Raf-dependent inhibition of MAPK/ERK kinase) (Banihashemi and Albert, 2002). In rat striatum and frontal cortex, D1Rs signal by G(s) to adenyl cyclase and by G(q) to the PI pathway (PI-4,5-diphosphate ! PLC). In contrast, in the amygdala and HC, D1Rs bind only to G(q) proteins (Jin et al., 2001). In transfected HEK293 cells, the structurally related D1 and D5 receptors use diVerent G-proteins composed of distinct subunits to stimulate adenyl cyclase (Wang et al., 2001). Regulators of G-protein signaling proteins (RGS) promote the return of G-proteins after

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activation to their inactive form. Various stimuli induce fast gene expression of RGS. In rat striatum, RGS2, 4, 8, 9, and 10 are the most abundant forms. Of these RGS2 regulates the D1 ! Golf ! adenylcyclase5 pathway, and RGS4 regulates the D2 ! Go ! adenylcyclase5 pathway (Taymans et al., 2003). In rat striatum, D1Rs upregulate and D2Rs downregulate RGS2 expression (Geurts et al., 2002). B. The ERK/MAPK and CREB Pathways In rat glioma cells, D2Rs possess two independent signaling pathways (a) an intracellular redox–kinase pathway leading to DNA synthesis and mitogenesis and (b) by means of p38 MAPK (mitogen-activated protein kinase) to induce glial fibrillary acidic protein (GFAP) expression (Luo et al., 1999). In rat brain slices, D2Rs activate two distinct signaling pathways (a) by means of increased i.c. Ca2þ levels ! PKC ! MAPKK ! MAPK and (b) by means of PKC ! Ca2þ/calmodulin–dependent protein kinase (CCPDK [a.k.a. CAMK]) ! cAMP response element–binding protein (CREB). Both these pathways need dopamine and cAMP-regulated phosphoprotein (DARPP)–32 activity (Yan et al., 1999). In rat HC CA1, several neuromodulatory neurotransmitter receptors (including DARs) couple to MAPK by means of PKA and/or PKC. This leads by phosphorylation of CREB to modulated gene expression (Roberson et al., 1999). In rat pituitary lactomorphs, D2Rs coupled to G(o) inhibit prolactin gene expression by means of ERK1/2 (Liu et al., 2002). The CREB protein plays a key role in DA-mediated nuclear signaling and neuroplasticity in the intact striatum but not in L-DOPA–treated 6-OH DA–lesioned rats. Here, the AP-1 transcription factor seems to take over this role (Andersson et al., 2001). D1R stimulation induces phosphorylation of CREB in the nucleus by means of NMDA receptor–mediated Ca2þ signaling (Dudman et al., 2003). Stimulation of D1Rs induces the phosphorylation of Ser897 on the NR1 subunit by protein kinase A (PKA) crucial for D1 receptor-mediated CREB phosphorylation. The authors conclude that this recruitment of the NMDA receptor signal transduction pathway by D1 receptors may provide a general mechanism for gene regulation that is fundamental for mechanisms of drug addiction and long-term memory (Dudman et al., 2003). In the mesolimbic dopamine reward pathway, the nucleus accumbens contains considerably higher levels of CREB than does the VTA, suggesting the greater importance of the former than the latter in CREB nuclear activities related to reward (Walters et al., 2003). In rat striatal slices, depolarization by Kþ activates extracellular signal– regulated protein kinase 1/2 (ERK1/2) and thus increases the phosphorylation of tyrosine hydroxylase at ser 19, ser 31, and ser 40. This upregulates the enzyme

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and increases DA synthesis (Lindgren et al., 2002). In contrast, D2Rs inhibit adenyl cyclase and so downregulate tyrosine hydroxylase and DA synthesis by reducing cAMP-induced phosphorylation at ser 40 (Lindgren et al., 2001). C. The cAMP Pathway In primary cultures of rat HC and cortex, Lezcano and Bergson (2002) report that there are two pathways of signaling induced by D1/D5R activation (a) by Ca2þ and (b) by cAMP. The striatum possess only pathway (b). DA autoreceptors inhibit DA release, DA synthesis, and firing of DA neurons. They can couple either to cAMP or to PI signaling chains. In mouse brain, autoreceptors couple preferentially to the former (Mercier et al., 2001).

D. More on the CAMKII Pathway In rat HC slices, D1Rs and D2Rs inhibit CAMKII activity (Hou and Zhang, 1999). In rat striatum, all four isoforms of CAMKII (, , , and ) are found. In NG108–15 cells transfected with D2LRs, activation of this receptor upregulated nuclear CAMKII, as well as BDNF, thereby mediating Ca2þ-dependent gene modulation (Takeuchi et al., 2002).

E. Further on Cascades Involving Phosphorylation Dopamine-regulated and cyclic AMP–regulated phosphoprotein of molecular weight 32,00 (DARPP-32) is concerned in a very large number of signaling systems and forms a gateway where many influences coincide, leading to many downstream processes. Greengard and co-workers (Svenningsson et al., 2004) have stressed the key role that DARPP-32 plays in neuronal signaling. The abstract of this important article states that it was identified initially as a major target for dopamine and protein kinase A (PKA) in striatum. However, recent advances now indicate that regulation of the state of DARPP-32 phosphorylation provides a mechanism for integrating information arriving at dopaminoceptive neurons, in multiple brain regions, by a variety of neurotransmitters, neuromodulators, neuropeptides, and steroid hormones. Activation of PKA or PKG stimulates DARPP-32 phosphorylation at Thr34 and thereby converts DARPP-32 into a potent inhibitor of protein phosphatase-1 (PP–1). DARPP-32 is also phosphorylated at Thr75 by Cdk5, and this converts

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DARPP-32 into an inhibitor of PKA. Thus, DARPP-32 has the unique property of being a dual-function protein, acting either as an inhibitor of PP-1 or of PKA. The state of phosphorylation of DARPP-32 at Thr34 depends on the phosphorylation state of two serine residues, Ser102 and Ser137, which are phosphorylated by CK2 and CK1, respectively. By virtue of its ability to modulate the activity of PP-1 and PKA, DARPP-32 is critically involved in regulating electrophysiological, transcriptional, and behavioral responses to physiological and pharmacological stimuli, including antidepressants, neuroleptics, and drugs of abuse (Svenningsson et al., 2004). The same authors (Svenningsson et al., 2004) also identify DARPP-32 as a common factor in the signaling chains activated by the three main classes of psychotomimetic drugs (i.e., those acting on the dopamine [amphetamine], serotonin [LSD], and glutamate [PCP] systems. In mouse dorsal striatum, D2SRs on presynaptic DA terminals participate in the downregulation of tyrosine hydroxylase phosphorylation. D2LRs are involved at postsynaptic sites on medium spiny neurons, where they modulate the phosphorylation of DARPP (Lindgren et al., 2003). Striatonigral projection neurons carry mainly D1Rs, whereas striatopallidal neurons carry mainly adenosine A2ARs and D2Rs. In striatal slices, D2Rs strongly inhibit the phosphorylation of DARPP-32 that is induced either by D1R or by A2AR activation. Thus, D2Rs decrease DARPP-32 phosphorylation in striatonigral and in striatopallidal projection neurons by two diVerent mechanisms (Lindskog et al., 1999). In mouse neostriatal slices, cyclin-dependent kinase 5 (Cdk5) increases phosphorylation of DARPP-32 at thr 75. D1Rs inhibit this response by way of PKA and so reverse the eVect of Cdk5. Thus, Cdk5 and PKA signaling at this locus are mutually antagonistic (Nishi et al., 2000). In rat neostriatum, D1Rs inhibit GABA(A) receptor currents by means of the cascade PKA!DARPP-32!PPI that targets the GABA(A)R -1 subunit (Flores-Hernandez et al., 2000). DA receptors use the inositol phosphate–signaling pathway extensively. In rat brain slices, D1Rs stimulate PI hydrolysis and thus the formation of two second messengers inositol triphosphate (PI3) and diacetyl glycerol (DAG). In a mouse HC-neuroblastoma hybrid cell line, D1Rs couple to PLC by a G(q) protein and also stimulate inositol phosphate synthesis (Jin et al., 1998). In mouse embryonic striatal neurons, D1Rs and D2Rs both activate the pathway PI3K!Akt!nuclear phosphorylation of transcription factors. Akt is a prosurvival ser/thr kinase that is activated in response to trophic factors (Brami-Cherrier et al., 2002). In rat striatal homogenates, D1RS also mediate the phosphorylation of striatal-enriched protein tyrosine phosphatase (STEP) by PKA (Paul et al., 2000). In neonatal hamster suprachiasmatic nucleus (SCN), DA promotes the phosphorylation of CREB in GABA cells by means of PKA and MAPK (but not CaMKII/IV). In this location, glu acts on a diVerent pathway by way of Ca2þ

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and CaMKII, which is responsible for the entrainment of the light/dark cycle (Schurov et al., 2002). Young and Yang (2004) show that depolarizing state– dependent, D1/D5–activated, bi-directional switching of soma-dendritic L-type Ca2þ channels by way of PKA-dependent potentiation and PKC-dependent suppression may provide spatiotemporal regulation of synaptic integration and plasticity in PFC.

V. Miscellaneous Items

A. The Cytoskeleton DA also aVects the actin cytoskeleton in cells. In the prolactin-secreting cells of the pituitary (lactomorphs), D2Rs induce actin assembly and stabilization that is linked to DA-stimulated inhibition of prolactin secretion. This is mediated by Ca2þ and, in some cells, by cAMP as well. This response also involves modification of the expression and localization of the actin-binding proteins talin and actin but not vinculin. In particular, DA promotes the association of talin to cell membranes, thus modulating cell shape (Nguyen et al., 1999). Lin et al. (2001) make the interesting observation that the proper cell surface location of D2Rs and D3Rs requires links between filamin A with the cytoskeleton. B. Reactive Oxygen Species Until recently, it was thought that reactive oxygen species (ROS) were merely unimportant by-products of the activity of certain brain enzymes such as cyclooxygenase. Now, however, it is apparent that some of these ROS, in particular hydrogen peroxide (H2O2), are second messengers in their own right (Smythies, 1997, 1999a, b). DA and ROS interact in various ways. In guinea-pig dorsal striatal slices, H2O2 inhibits synaptic DA release by means of a Ca2þ-dependent mechanism (Chen et al., 2001). In rat HC, ROS inhibit the DA, but not the NE, transporter (Haughey et al., 1999). In rat astroglial cultures, application of DA leads to increased superoxide production accompanied by the oxidation of glutathione (GSH) to GSSH. It also leads to increased production of H2O2, which results in the eZux of GSSG from the cell (Hirrlinger et al., 2002). In cultured human neurons, DA induces apoptosis by causing a decrease in the mitochondrial membrane potential mediated by the production of excess ROS (Medina et al., 2002). Malonate is a neurotoxin that induces striatal lesions similar to those seen in Huntington’s disease. This is brought about by two independent

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mechanisms (1) DAT-dependent (and DR-independent) production of ROS that leads to depletion of i.c. DA stores, and (2) excessive stimulation of D2Rs (Xia et al., 2001). ROS such as peroxynitrate can oxidize DA to form toxic DA quinones (Smythies, 1999). DA can also play a neuroprotective role and inhibits ROS-induced cell death. It does this partly because it is itself an antioxidant and free radical scavenger (Liu and Mori, 1993) and partly by means of D2R-mediated stimulation of the synthesis of i.c. antioxidant enzymes (Sawada et al., 1998; Smythies, 1999a, b). D4Rs inhibit ROS-induced cell death. This is mediated by blockade of the opening of the cGMP-operated Ca2þ channel that constitutes a late phase in glu-induced cell death (Ishige et al., 2001). In rat neostriatum, DA innervation normally suppresses OH formation, whereas l-DOPA has little eVect (Kostrzewa et al., 2000). The antioxidant carnosine, which is co-released together with glu from glu terminals, protects neurons against methamphetamine-induced toxicity evidenced by increased heat shock protein P27 production (a measure of astroglial stress) but fails to protect against methamphetamine-induced loss of DA terminals (Pubill et al., 2002). C. Interactions between Dopamine and Arachidonic Acid Arachidonic acid (AA) is an important volume-signaling agent produced by the action of PLAII on membrane lipids. In mouse striatum, AA increases the DA release evoked by NMDA and carbachol. This eVect is partly Ca2þ dependent (L’hirondel et al., 1999). In rat striatal synaptosomes, AA also increases DA release and inhibits DA uptake by PKC (L’hirondel et al., 1995). In goldfish pituitary cells, AA mediates growth hormone release by D1Rs that act distal to cAMP formation. Calmodulin is also involved in this response, acting downstream from cAMP and PKC (Chang et al., 1996). In transfected CHO cells, D2Rs increase the AA release triggered either by purines or by an increase in i.c. Ca2þ levels. D1Rs by themselves have no such eVect, but they can act synergistically with D2Rs to increase AA release (Piomelli et al., 1991) acting by means of PLA2 (Vial and Piomelli, 1995). D. Endocytosis of Dopamine Receptors Most receptors for neurotransmitters and neuromodulators are endocytosed after stimulation and are transported to the endosome system (Smythies, 2004). DRs are no exception. D1Rs in rat striatum cell dendrites and somata are located mainly extrasynaptically. On stimulation by DA or a DA agonist,

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they endocytose and translocate to endosomes (Dumartin et al., 1998). In transfected CMO cells, D2RS, on stimulation, are endocytosed when the receptor is in its low-aYnity state (Ko et al., 2002). In COS–7 and HEK–293 cells, activation of D2Rs induces a G-protein–coupled kinase-dependent phosphorylation of the D2R receptor that is followed by a robust translocation of the endocytosis-associated molecule -arrestin to the cell surface, leading in turn to massive receptor internalization. This process is mediated by a clathrin-coated pathway in which dynamin is involved. The D2R is then transported to a diVerent endosome from that to which endocytosed NE -adrenergic receptors are transported. In the case of the D3R, in contrast, only a minor degree of internalization occurs (Kim et al., 2001). Activation of the D4R in the rodent PFC not only inhibited NMDAR-induced currents but also triggered endocytosis of the NMDARs. In contrast, D1Rs inhibit endocytosis of NMDARs (Wang et al., 2003). Dopamine-releasing protein (DARP) is a potent regulator of DA release that is involved in the development of the catecholinergic system. Smith and Ramirez (2002) studied the endocytosis of the N-terminal sequence of DARP in rat C6 glioma cells and mesencephalic cell cultures. Electron microscopy revealed that the peptide is rapidly endocytosed and transported to multivesicular bodies, endosomes, and lysosomes in a clathrin-dependent manner. Tyrosine kinase activity may also be involved (Lin et al., 2001). E. Volume and Synaptic Transmission As noted previously, much DA transmission in some areas is mediated outside the synapse by volume transmission. In this process, DA-releasing varicosities simply release DA into the extracellular space of the neuropil, where it diVuses to excite any DA receptors that may happen to be in the vicinity. In rat basal ganglia, there is a large component of nonsynaptic DA varicosities (Yung et al., 1995). Caille et al. (1996) found abundant and widespread distribution of D1Rs at extrasynaptic sites in rat striatum and SN. Descarries et al. (1996) reported in rat striatum that 60 to 70% of DA varicosities were extrasynaptic and 30 to 40% were synaptic. The latter were always symmetrical, and 67% were located on dendrites, 30% on spines, and only 3% on somata. Smiley et al. (1994) found in primate cortex that D1Rs were mainly either nonsynaptic or were attached to the side of dendritic spines. In guinea-pig brain slices, volume transmission in the midbrain is regulated by DA diVusion, whereas in the striatum it is regulated by DA uptake (Cragg et al., 2001). In the striatum, the DA transporter (DAT) is located mainly on DA axons and terminals outside recognized synapses. In the SN, in contrast, the DAT is

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located specifically on dendrites and on presynaptic and postsynaptic zones, indicating that some form of synaptic transmission is important in this location (Hersch et al., 1997; but vide Caille et al., 1996). In monkey N.Ac. (medial subdivision), most DA contacts are synaptic, with 50% on dendritic shafts, 44.2% on spines, and only 5.1% on somata (Ikemoto et al., 1996). In rat locus ceruleus, there is a dense DA innervation with many axospinous synapses on NE neurons (Maeda et al., 1994). A good general review of volume transmission is given by Zoli et al. (1998). A review of the possible role of reversed transport of DA by DAT in volume transmission is also given. DA dendrodendritic release entails reversal of the DAT rather than conventional exocytosis (Falkenburger et al., 2001). There are marked diVerences in the cellular control of dendrodendritic release of DA in the striatum and the SNpc, with a minimum role for Ca2þ in the latter (Chen and Rice, 2001). On the basis of rat experiments, Castagne et al. (2004) suggest that low brain glutathione and ascorbic acid levels associated with a perturbation of the dopaminergic system actively participate in the development of some cognitive deficits aVecting schizophrenic patients.

F. Back Propagation The action potential fired by a neuron does not only go down the axon but also back propagates into the dendritic tree. This process is important for Hebbian learning. In HC CA1 pyramidal cells, back propagated dendritic potential attenuates the further they get along the progressively thinning apical dendrite. This attenuation is inhibited strongly by activation of -adrenergic and muscarinic acetylcholine receptors and only in a subpopulation by dopamine receptors (HoVman and Johnston, 1999). The authors suggest that this diVerential result between these three transmitter systems may reflect the number of their terminals in this locus. The fact that these three neuromodulators can enhance back propagation in distal dendrites has important consequences for the role they might play in Hebbian learning and related mechanisms.

G. Tropins Dopamine interacts with various tropins in the brain. In mice, D2R antagonists such as haloperidol increase nerve growth factor (NGF) gene expression. This is mediated by c-fos signaling to the AP-1 binding site on the NGF gene. This leads to ACh release (Ozaki, 2000).

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In PC12 cells, brief treatment with NGF induces depolarization, and an increase in ionomycin-stimulated DA and ACh release. This is mediated by raising the number of Ca2þþ-responsive secretory vesicles in the axon terminal. PI3-K and MAPK pathways are essential for this eVect (Amino et al., 2002). In postnatal mouse striatum, BDNF inhibits DA-triggered oxidative stress and partly blocks basal and DA-induced apoptosis. BDNF also increases DARPP-32 expression (Petersen et al., 2001). In rat striatal slices and synaptosomes, BDNF increases the synaptic output of DA by means of PI–3K and Ras MEK (but not PLC ) pathways (Goggi et al., 2003). D1Rs increase levels of glial cell line–derived neurotrophic factor (GDNF) in human fetal astrocytes (Kinor et al., 2001). In rat SNpc, GDNF increases the expression of the cell adhesion molecules integrin and NCAM, leading to increased DA neuron survival, outgrowth, and DNA turnover (Chao et al., 2003). In rat postnatal DA neurons in culture BDNF, GDNF and cAMP mediate convergent signals, leading to increased survival of the DA neurons (Lara et al., 2003). H. Synaptic Plasticity Dopamine aVects various aspects of the many mechanisms that control synaptic plasticity. Some of these have been touched on previously. In the rat HC, D1Rs upregulate the cAMP pathway that increases the expression of the immediate early gene c-fos involved in synaptic plasticity (Kang et al., 2000). Berke et al. (1998) list more than 30 genes whose expression is rapidly increased by D1R activation. These include many involved in cellular and synaptic plasticity. In rat striatum, D2Rs block immediate gene expression but not by way of PKA (Adams and Keefe, 2001). D1Rs and D2Rs increase synaptic density in the rat PFC but not in parietal cortex. Conversely, the 5-HT depleter cCPA resulted in a decrease of 5-HT in the parietal cortex but not in the PFC (Sugahara and Shiraishi, 1998). In rat N.Ac., DA denervation by injection of 6-OH DA into the medial forebrain bundle led to the development in the core of shorter and less spiny dendrites and in the shell of less spiny dendrites (Meredith et al., 1995). In rat striatum, D1Rs induce a massive upregulation of the mRNAs for the synaptic proteins synaptotagmin I and IV (Glavan et al., 2000). DA also aVects another synaptic protein by promoting SNARE complex formation out of its component parts synaptobrevin, syntaxin, and SNAP25 (Fisher and Braun, 2000). Estrogen stimulation of neurite growth in mouse midbrain DA neurons depends on cAMP ! PKA signaling (Beyer and Karolczak, 200). A general review of the role of neurotransmitters in the cortex in cortical plasticity is given by Gu (2002).

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I. Dopamine and Neuronal Development Neurotransmitters, including dopamine, play a role in guiding the development and location of neurons. In mouse developing cortex, D1Rs upregulate dendritic growth by phosphorylating microtubule-associated protein-2 (MAP-2) by means of PKA in two steps. During the first 24 hours after stimulation, there is an initial weak response followed (24 to 48 hours) by a much more robust late response (Song et al., 2002). In the human neuronal NMB line, DA has a biphasic eVect on cell growth and diVerentiation. At low doses (20 to 50 M), DA decreases cell growth, induces an apparently diVerentiated cell morphology, and increases DA uptake. Higher doses (100 to 300 M) were neurotoxic and induced apoptosis. Further tests revealed that DATs were involved in both cases (Porat et al., 2001). In rat somatosensory cortex, the D3R is selectively and transiently expressed in the barrel neurons between the first and second postnatal weeks. This event is necessary for the barrel-like pattern to develop. After a lesion of the ventrobasal nucleus of the thalamus, D3Rs and D3 mRNA levels are much reduced in somatosensory cortex, and the barrels do not develop. Thus, an intact thalamocortical input is essential for D3Rs to occur in somatosensory cortex (Gurevich et al., 2001). The protein reelin controls neuronal migration and positioning and so plays a key role in brain development. Reelin mutations lead to a widespread disruption of laminar cortical development and proper development of subcortical structures. These include the DA neurons in the SNpc that fail to migrate properly from the origin in the tegmental area. This reelin may be supplied from the axons of striatal neurons (Nishikawa et al., 2003). Maternal deprivation in weanling rats upregulates and in neonatal rats downregulates DA function. The latter is accompanied paradoxically by increased release of DA in response to amphetamine and Kþ (Hall et al., 1999). The transcription factors En1, Pitx3, Nurr1, and Lmx1b are all important for the development of the mesencephalic dopaminergic system (Smidt et al., 2003). Neither nitric oxide nor lipid peroxidation are involved in the normal processes, including apoptosis, that occur during development of the SNpc (Groc et al., 2002).

VI. Dopamine and Learning

A. Long-Term Potentiation Learning in the brain is often studied by a phenomenon known as longterm potentiation (LTP). In this, a tetanic stimulus is applied to the input that increases the eYciency of the synapse. Long-term depression (LTD) is when a

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low-frequency stimulus decreases the eYciency of the synapse. DA plays a prominent role in this system, particularly in the PFC that receives DA input from the VTA, as well as an excitatory input from the HC. Both these pathways target in close proximity to each other dendritic spines of the pyramidal cells in layers V to VI. Activation of the DA pathway induces a long-lasting potentiation of the LTP produced by stimulation of the HC–PFC pathway (Gurden et al., 1999). In the rat PFC, D1Rs (but not D2Rs) potentiate NMDA-dependent LTP at the synapses made by axons from the HC by means of cAMP and PKA (Gurden et al., 2000). In rat dentate gyrus, D1/D5Rs induce LTP in vivo (Kusuki et al., 1997). In rat HC CA1, D1/D5Rs increase the magnitude of early LTP by means of cAMP as a second messenger and possibly by way of NMDA channels (Otmakhova and Lisman, 1996). These authors note that cAMP-dependent PKA is coanchored with protein phosphatase 2B and PKC in the postsynaptic density and dendritic cytoskeletal elements by a common anchoring protein, Furthermore, cAMP levels in neurons are modulated by i.c. Ca2þ levels, various G-proteins (G(s), G(i)), and the / subunit), PKC, and by the degree of membrane depolarization. Maintenance of the late stage of LTP in rat HC slices requires increased protein synthesis and D1R-mediated glycoprotein fucosylation (Angenstein et al., 1992). D1Rs selectively facilitate the expression of ischemia-induced LTP on the AMPA- (but not the NMDA-) mediated component of EPSPs of corticostriatal synaptic potentials (Saulle et al., 2002). B. LTP/LTD DA can promote LTP or LTD, depending on circumstances. In rat PFC layer V pyramidal cells, tetanization of aVerents from cortical layers I to III in the absence of DA resulted in either LTP, LTD, or no eVect in diVerent cells. The same stimulus in the presence of DA now led to LTD in most cells and LTP in none (Law-Tho et al., 1995). In rat PFC slices, Blond et al. (2002) found that DA can induce LTP or LTD according to circumstances; 50 Hz tetanization induced LTP in the slice if coupled in real time with the presence of DA in the bath. However, if the slice was exposed to DA 30 minutes before the tetanization, LTP resulted. D2R-null mice show severe locomotor impairment. In these mice, tetanic stimulation of corticostriatal fibers leads to LTP (NMDARdependent), whereas in normal mice, it leads to LTD not NMDAR-dependent (Calabresi et al., 1997). In the rat HC, CA1 D1/D5Rs potentiate LTP and inhibit LTD, whereas in rat dentate gyrus, these receptors have no eVect on LTP but still inhibit LTD (Kulla and Manahan-Vaughan, 2000). In rat HC granule cell layer, D2Rs block basal synaptic transmission and also inhibit LTD but have little eVect on LTP (Manahan-Vaughan and Kulla, 2003). In mouse midbrain slices, DA

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blocks the induction of LTD but has minimal eVect on LTD in N.Ac. (Thomas et al., 2000). In eNOS-deficient mice, NO promotes LTP and reduces LTD (Doreulee et al., 2003). In rodent striatum, D1Rs and D2Rs act synergistically to allow LTD formation, but they operate in opposition during the induction phase of LTP. Corticostriatal synaptic plasticity depends on an intact DA system and requires DARPP-32. The formation of LTD requires PKG, and of LTP requires PKA activity (Calabresi et al., 2000; Centonze et al., 2001). Depotentiation of LTP is the reversal of LTP by low-frequency stimulation or by anoxia. In rat HC, CA1 D1/D5Rs enhance LTP and also inhibit depotentiation by a cAMP-dependent and PKA-dependent mechanism. LTD requires the signaling chain calcineurin ! activation of PPI ! dephosphorylation of CAMKII, which acts as a switch on the AMPAR channel. This chain is inhibited by cAMP (Otmakhova and Lisman, 1998). In rat striatum, electrical stimulation of the cortex induces LTD in the corticostriatal input. In intact, but not in DA-depleted animals, this was prevented or reversed by concomitant stimulation of the SN (Reynolds and Wickens, 2000).

C. Miscellaneous DA eZux in the PFC is increased in a phasic manner when a rat engages in search behavior for food reward (Phillips et al., 2004). These researchers demonstrate a close functional relationship between the release of DA from terminals within the PFC and the retrieval of specific trial unique memories; furthermore, the magnitude of mesocortical DA eZux is predictive of the accuracy of this form of memory (Phillips et al., 2004). Roitman et al. (2004) measured the release of dopamine in N.Ac. every 100 msec during a conditioned lever press for sucrose reward. They found that dopamine release occurred 0.2 sec after the conditioned stimulus was presented and thereafter subsided. They concluded that dopamine release represents a learned association between the cues and sucrose availability.

VII. Dopamine and Behavior

A. Appetitive Learning DA neurons in the SN and VTA do not simply respond to rewarding stimuli as was once thought but respond to salient stimuli whether novel, intense, rewarding, aversive, or conditioned rewarding (Horvitz, 2002). In particular, any unexpected change in the environment causes the DA neurons to fire. The

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resulting increase in DA levels in the striatum (dorsal from SN, ventral from VTA) gates the input from the sensory and motor cortex and, particularly, the input from two areas of brain—orbitofrontal cortex and basolateral amygdala— that are concerned with incentive/motivational reward signaling. The DA so released promotes strong glu currents and inhibits weak ones. Thus, the role of DA in appetitive conditioning is not to tell the striatal cells that a reward has been received—the input from the orbitofrontal cortex and basolateral amygdala does that—but to convey the information that an important event has occurred in the environment (Horvitz, 2002). Legault and Wise (2001) confirm that a novel stimulus in freely moving cats leads to increased DA release in N.Ac. (part of the ventral striatum) and increased exploratory behavior. Neurons in the thalamic centromedian/parafascicular complex also respond to unexpected stimuli. This complex has a massive two-way projection to and from the striatum. Thus, these key midline thalamic nuclei can also provide striatal neurons with significant information that can activate conditioned responses in combination with DA inputs (Matsumoto et al., 2001). According to Berridge and Robinson (1998), the role of dopamine in the brain is to contribute ‘‘incentive salience attributions’’ to stimuli (i.e., that DA is necessary for ‘‘wanting,’’ not for ‘‘hedonic import’’ or for ‘‘associated reward learning’’). The PFC influences the behavioral impact of activation of the basolateral amygdala by means of a concomitant suppression of DA release in N.Ac. in a triadic relationship. In freely moving rats, microstimulation of the basolateral amygdala, at intensities that produce a mild behavioral activation, evoke a rapid increase in glu eZux in the PFC and N.Ac. shell. This is accompanied by increased DA release in the PFC but not in the N.Ac. shell. An increase in DA release in N.Ac., during this period of stimulation, was seen only if glu activation in the PFC was inhibited at either the presynaptic or postsynaptic level. When the PFC fails to inhibit the release of DA in N.Ac., aberrant behavior (abnormal perseveration of a response after the stimulus has stopped) results after stimulation of N.AC. (Jackson and Moghaddam, 2001). In rat N.Ac., coactivation of NMDARs (corticolimbic) and D1Rs (from the VTA) in the core area is a key process in the acquisition of instrumental learning (Smith-Roe and Kelley, 2000). In cat sensorimotor cortical pyramidal cells, D1RS and D2Rs facilitate conditioned reflex formation (Storozhuk et al., 2002). In rat brain, the medial PFC attenuates sensory-driven amygdala-mediated aVective responses by recruitment of inhibitory neurons in the basolateral amygdala, which suppress sensory cortical inputs. In basolateral amygdala (BLA), input from the temporal cortex drives BLA output neuron firing, whereas input from the medial PFC drives the firing of inhibitory interneurons. Normal rises in DA levels in the BLA may facilitate appropriate aVective responses, but pathological increases in DA levels in this nucleus produced by drugs (cocaine, amphetamine), stress, or disease (schizophrenia) dampens this PFC-evoked attenuation, leading to potentially

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disruptive disinhibition of sensory-driven aVective responses (Rosenkranz and Grace, 2001). Amygdala nuclei also interact with the mesocorticolimbic DA system during food reward. Inactivation of the central (but not the basolateral) nucleus by local injection of lidocaine impairs the development of satiety during eating. This is associated with decreased basal eZux of DA before feeding and an attenuated increase in DA eZux related to anticipatory and consummatory aspects of feeding behavior in both N.Ac. and PFC. This interaction between the central nucleus of the amygdala and the DA system may represent a mechanism by which hunger and satiety signals influence the estimation of the value of a food reward. Alternatively, it may represent a mechanism by which memory for a recently consumed meal modulates food intake (Ahn and Phillips, 2002). In the rat N.Ac., eating evokes D2R-dependent inhibition of glycine release (Saul’skaya et al., 2001). In rat globus pallidus, eating evokes a 130% increase in e.c. DA levels (Hauber and Fuchs, 2000). Akirav and Richter-Levin (2002) give an excellent review of the role of the basolateral amygdala in the complex modulation of hippocampal plasticity. Chronic stress in the rat induces impairment in working memory for a T-maze task. This is associated with a marked reduction in DA transmission in PFC plus a possibly compensatory increase in D1R density (Mizoguchi et al., 2000). In a number of species (cat, rat, mouse, monkey), D4Rs modulate exploratory behavior by action in striatal striosomes rather than in striatal matrix or in N.Ac. In contrast, D4Rs modulate motor coordination by action in the striatal matrix (Riviera et al., 2002). PET studies in humans show an inverse correlation between a low threshold for cold pain and high D2 binding levels in the putamen, whereas there was an inverse relationship between cold pain tolerance and D2 binding in the right medial temporal cortex. The magnitude of heat pain threshold elevation produced by concurrent cold pain was directly correlated with D2 binding in the left putamen (Hagelberg et al., 2002). Fear conditioning involves the induction of LTP of excitatory synaptic transmission in the lateral amygdala. DA gates this by suppressing feed forward inhibition from local INs (Bissiere et al., 2003). In the monkey PFC layer V pyramidal cells, DA, by a presynaptic action, inhibits local horizontal excitatory transmission that is part of the network that sustains local memory. This correlates with the finding by Romanides et al. (1999) that increased DA tone (or reduced glu tone) in the rat PFC disrupts working memory. The dopamine system is also involved in attention, where it modulates its more ‘‘executive’’ aspects, such as attentional set-shifting and working memory. In contrast, NE and ACh are involved in ‘‘low level’’ aspects of attention such as attentional orienting (Coull, 1998).

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B. Prepulse Inhibition D1Rs, but not D2Rs, reduce prepulse inhibition of the startle response in rat in HC CA1 neurons (Ellenbroek et al., 2002). In rat basolateral amygdala, local infusion of NMDA and GABA(A) antagonists reduces prepulse inhibition. This eVect was reversed by the DA antagonist haloperidol (Fendt et al., 2000).

VIII. Parkinson’s Disease

Pathology of the brain’s dopamine system plays a key role in the genesis of this disease. The main finding is a progressive destruction of neuromelanincontaining neurons. These can be dopaminergic, as in the SNpc and VTA, or noradrenergic, as in the locus ceruleus and medullary A1-A3 group. Neuromelanin is composed mainly of a complex polymer of 5,6-dihydroxyindole, and possibly 5-cysteinyldopamine, on a glycoprotein matrix. 5-Hydroxyindole is the end product of dopamine and NE oxidation pathways in which these catecholamines are converted sequentially into various quinones, including the highly neurotoxic o-semiquinone. Normally, neuromelanin is neuroprotective because it is a potent antioxidant, mops up toxic catecholamine o-quinones, and chelates large amounts of toxic heavy metals. However, in excess, it becomes neurotoxic, largely by disruption of the cell by large amounts of the dense polymer. In the SNpc in Parkinson’s disease, there is evidence of severe oxidative stress and excessive production of dopamine o-quinones. There is excessive oxidation of proteins in SNpc (and cortex) (Castellani et al., 1996; Jenner and Olanow, 1998), excess products of lipid peroxidation in the SNpc (Nakamura et al., 1997), mitochondrial defects (Itoh et al., 1996; Schapira et al., 1992) probably caused by DA quinone toxicity (Zhang et al., 1998) and low glutathione (GSH) levels (Merad-Boudia et al., 1998; Nakamura et al., 1997; Pearce et al., 1997), excess dopamine o-quinone synthesis (Mattammal et al., 1995), and excess levels of 5-cysteinyldopamine in the CSF, which also indicates excess dopamine o-quinone synthesis, because the latter are required steps in the synthesis of the former (Cheng et al., 1996). Shen and Dryhurst (1996) also found low GSH levels in the SNpc but normal levels of its oxidized product GSSG. This suggested that GSH is depleted by forming 5-cysteinyldopamine and, thus, scavenging dopamine o-quinones rather that by its more usual role of scavenging ROS. This finding is unique to the SNpc in PD and is not found in other neurodegenerative diseases. In DA neurons in PD, COX–2 activity is upregulated. This enzyme not only produces ROS but also oxidizes DA to DA quinones (Teismann et al., 2003). MAPT toxicity acts by way of COX–2 (Feng et al., 2002). Thus, Parkinson’s disease may be characterized by low levels of antioxidant defenses in the aVected

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brain nuclei plus high levels of catecholamine oxidation and oxidative stress (Smythies, 1999a, b). The pattern of loss of SN neurons follows a fixed pattern. It starts in nigrosomes and later involves the matrix. The origin is in the main pocket nigrosomes 1 in the caudal and mediolateral part of the SNpc and then spreads to nigrosomes 2 ! 4 ! 3 ! 5 (a medial and rostral progression). The matrix shows a similar, but less well marked, progression (Damier et al., 1999). Dramatic neuron loss also occurs in PD (and in progressive subnuclear palsy) in the caudal intralaminar nucleus of the thalamus, but not in any other thalamic nuclei (Henderson et al., 2000). Loss of DA neurons in PD provokes a change in corticostriatal synapses. Fifty percent of these become perforated synapses (very active) located on spines that carry D1Rs. There is no such change on spines that do not carry D1R (Muriel et al., 2001). The authors suggest that this may represent a compensatory hyperactivity in response to loss of DA innervation. DA denervation of the striatum (by 6-OH DA) leads to modulation of 50 genes, including those involved in the intracellular transduction of the DA signal and in the regulation of glu transmission (Napolitano et al., 2002). Mice with D2R gene knockout develop a PD-like syndrome by way of basal ganglia dysfunction (Fowler et al., 2002). Oxidative stress leads to the death of SN, DA cells being a primary pathogenic mechanism in PD. This involves mitochondrial complex 1 malfunction. It is not surprising, therefore, that oxidative stress modulates the expression of several groups of genes in DA cells, including those for functional molecules in mitochondrial complex 1 function, exocytosis, traYcking of membrane proteins, oxidoreductase, molecules that modulate apoptosis, and, in particular, the following genes the physiological dysfunction of whose products have been implicated in PD—mitochondrial complexes 1, B8, B19 (reduced) and syntaxin 8 and hemeoxygenase-1 (increased) (Yoo et al., 2003). Mutations of the -synuclein gene have been linked to rare autosomal dominant forms of PD. These mutations lead to impaired DA storage in vesicles (where DA is protected from oxidative attack) and so to increased DA oxidation in the cytoplasm (where DA is not so protected) (Lotharius and Brundin, 2002). In dopaminergic SH-SY5Y cells engineered to overexpress the A53T and A30P mutant forms of synuclein, such overexpression leads to raised levels of ROS and increases the susceptibility of the cell to DA-induced toxicity (Junn and Mouradian, 2002). In these cells, inhibition of the protease caspase protects DA neurons from 6-OH–induced cell death, yet neurons may remain functionally impaired (von Coelln et al., 2001). Low doses of the insecticide rotenone acts synergistically with the inflammatogen lipopolysaccharide (LPS) to induce degeneration of DA neurons. This is mediated by generation of ROS (superoxide) by brain microglial NADPH oxidase. This may be relevant to the multifactorial genesis of PD and the role of environmental toxins therein. Jenner (2003) has given a good review of the role of oxidative stress in P.D.

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IX. Miscellaneous Items

D5R-null mice have increased sympathetic tone and raised blood pressure. This is associated with oxytocin-dependent sensitization of vasopressin and nonNMDAR glu–mediated pathways (Hollon et al., 2002). The genes for DAT (SLC3A6) and the D4R are involved in attention deficit disorder (Di Maio et al., 2003). In rat striatum, iron deficiency leads to increased extracellular DA levels caused by depressed DAT function, not by increased DA release (Erikson et al., 2000). In rat striatum, perfusion of ferric chloride leads to DA release without immediate destruction of DA terminals (Prikhojan et al., 2002). A. Schizophrenia The role of dopamine in schizophrenia is a vast subject that is too extensive to be included in this review. Moreover, interest in schizophrenia research has now shifted away from possible abnormalities in individual transmitters, such as dopamine and glutamate, to disorders in the field of synaptic plasticity (for details see Smythies [2004]).

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Further Reading

Arbuthnott, G. W., Ingham, C. A., and Wickens, J. R. (2000). Dopamine and synaptic plasticity in the neostriatum. J. Anat. 196, 587–596. Beyer, C., and Karolczak, M. (2000). Estrogenic stimulation of neurite growth in midbrain dopaminergic neurons depends on cAMP/protein kinase A signalling. J. Neurosci. Res. 59, 107–116.

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Nitsch, C., and Riesenberg, R. (1995). Synaptic reorganisation in the rat striatum after dopaminergic deaVerentation: An ultrastructural study using glutamate decarboxylase immunocytochemistry. Synapse 19, 247–263. Otani, S., Blond, O., Desce, J. M., and Crepel, F. (1998). Dopamine facilitates long-term depression of glutamatergic transmission in rat prefrontal cortex. Neuroscience 85, 669–676. Reyes, M. G., Faraldi, F., Rydman, R., and Wang, C. C. (2003). Decreased nigral neuromelanin in Alzheimer’s disease. Neurol. Res. 25, 179–182. Rivera, A., Cuellar, B., Giron, F. J., Grandy, D. K., de la Calle, A., and Moratalla, R. (2002). Dopamine D4 receptors are heterogeneously distributed in the striosomes/matrix compartments of the striatum. J. Neurochem. 80, 219–229. Saal, D., Dong, Y., Bonci, A., and Malenka, R. C. (2003). Drugs of abuse and stress trigger a common synaptic adaptation in dopamine neurons. Neuron 37, 577–582. Sulzer, D., Joyce, M. P., Lin, L., Geldwert, D., Haber, S. N., Hattori, T., and Rayport, S. (1998). Dopamine neurons make glutamatergic synapses in vitro. J. Neurosci. 18, 4588–4602. Svenningsson, P., Tzavara, E. T., Carruthers, R., RachleV, I., Wattler, S., Nehls, M., McKinzie, D. L., Fienberg, A. A., Nomikos, G. G., and Greengard, P. (2003). Diverse psychotomimetics act through a common signaling pathway. Science 302, 1412–1415. Svensson, E., Wikstrom, M. A., Hill, R. H., and Grillner, S. (2003a). Endogenous and exogenous dopamine presynaptically inhibits glutamatergic reticulospinal transmission via an action of D2-receptors on N-type Ca2þ channels. Eur. J. Neurosci. 17, 447–454. Tessitore, A., Hariri, A. R., Fera, F., Smith, W. G., Chase, T. N., Hyde, T. M., Weinberger, D. R., and Mattay, V. S. (2002). Dopamine modulates the response of the human amygdala: A study in Parkinson’s disease. J. Neurosci. 22, 9099–9103. Tong, J., Ross, B. M., Sherwin, A. L., and Kish, S. J. (2001). Dopamine D1-stimulated adenylyl cyclase activity in cerebral cortex of autopsied human brain. Neurochem. Int. 39, 117–125. Umegaki, H., Munoz, J., Meyer, R. C., Spangler, E. L., Yoshimura, J., Ikari, H., Iguchi, A., and Ingram, D. K. (2001). Involvement of dopamine D(2) receptors in complex maze learning and acetylcholine release in ventral hippocampus of rats. Neuroscience 103, 27–33. Undie, A. S. (1999). Relationship between dopamine agonist stimulation of inositol phosphate formation and cytidine diphosphate-diacylglycerol accumulation in brain slices. Brain Res. 816, 286–294. Wang, X., Zhong, P., and Yan, Z. (2002a). Dopamine D4 receptors modulate GABAergic signaling in pyramidal neurons of prefrontal cortex. J. Neurosci. 22, 9185–9193. Wang, Z., Feng, X. Q., and Zheng, P. (2002b). Activation of presynaptic D1 dopamine receptors by dopamine increases the frequency of spontaneous excitatory postsynaptic currents through protein kinase A and protein kinase C in pyramidal cells of rat prelimbic cortex. Neuroscience 112, 499–508. Williams, S. M., and Goldman-Rakic, P. S. (1998). Widespread origin of the primate mesofrontal dopamine system. Cereb. Cortex. 8, 321–345. Wu, M., Hajszan, T., Leranth, C., and Alreja, M. (2003). Nicotine recruits a local glutamatergic circuit to excite septohippocampal GABAergic neurons. Eur. J. Neurosci. 18, 1155–1168. Yasuno, F., Suhara, T., Sudo, Y., Yamamoto, M., Inoue, M., Okubo, Y., and Suzuki, K. (2001). Relation among dopamine D(2) receptor binding, obesity and personality in normal human subjects. Neurosci. Lett. 300, 59–61.

SECTION III THE NOREPINEPHRINE SYSTEM

I. Microanatomy II. Functional Neuroanatomy A. Excitatory Effects B. Inhibitory Effects C. Biphasic Effects III. Electrical Activity A. Potassium Channels B. Calcium Channels IV. Interactions with Other Transmitters A. Norepinephrine and Glutamate B. Norepinephrine and GABA V. Postsynaptic Cascades A. cAMP B. MAPK/ERK C. Calcineurin D. Phosphatidylinositol E. Prostaglandins F. Nitric Oxide G. Reactive Oxygen Species H. Endocytosis I. Synaptic Plasticity J. Genes VI. Tropins and Cytokines A. Tropins B. Cytokines VII. Memory and Learning (LTP) A. Neuronal Development B. Stress C. Behavior VIII. Miscellaneous Items A. Alzheimer’s Disease B. Anticholinesterase Neurotoxicity C. Gap Junctions D. Oxytocin (OT)/Vasopressin (VP) E. Opioids F. Transporters G. Histamine References


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I. Microanatomy

The norepinephrine (NE)-containing neurons are located in the brainstem in the locus ceruleus (LC) and the adjacent A5 and A7 areas (Sawamura et al., 2000). The axons of these neurons conform to the usual pattern for neuromodulators of diffuse and sparse projections to all areas with little overlap. In some places, NE axons form terminal synapses with their target neurons, but in other places, volume transmission is predominant from boutons-en-passage that do not form specific synapses but release NE in a paracrine manner. For example, in the primate forebrain, NE axons form a bed of fine varicose axons that are codistributed with the cholinergic neurons of the nucleus basalis (Smiley et al., 1999). DA axons are 10 times more numerous throughout the primate basal forebrain than are NE axons, except in the medial septum, where the reverse is the case. 5-Hydroxytryptamine (5-HT) axons also form a dense plexus throughout the basal forebrain (Smiley et al., 1999). These three types of axons in the nucleus basalis end in boutons termineaux that synapse on cholinergic cells. In contrast, in the rat hippocampus (HC) CA1 stratum radiatum, many NE, 5-HT, and acetylcholine (ACh) axons carry numerous boutons-en-passage and few boutons termineaux on spines or dendrites, indicating volume transmission (Umbriaco et al., 1995). Likewise, in the primate dorsolateral prefrontal cortex (PFC), most boutons are en-passage and are not boutons termineaux (i.e., they are not in direct apposition with the 2 ARs that are scattered along the dendrites and axons of neurons and on astrocytes; Aoki et al., 1998). In this location, 2 ARs are particularly dense at presynaptic sites on axons. This suggests that most NE activity in the cortex is either mediated by volume transmission or by presynaptic activity (Aoki et al., 1998). In mouse cerebellar cortex, the fine beaded NE axons exhibit many boutons-en-passage along their lengths, none of which make synaptic contacts with the target neuronal elements (Abbott and Sotelo, 2000). Most of them were, however, close to the dendrites of Purkinje, granule, stellate, and basket cells without forming discernible synapses. This, again, is evidence for volume transmission in the cerebellar cortex. However, Milner et al. (2000) found in rat dentate gyrus that ARs are located ‘‘near’’ postsynaptic densities (PSDs) on the large dendrites and spines of granule cells. ARs are also located on astrocytes in this location in close proximity to presynaptic excitatory terminals on spines. Some ARs were also found on presynaptic unmyelinated axon terminals (Milner et al., 2000). In rat visual cortex, 2A ARs are found particularly in layers II to IV on proximal and distal dendrites, as well as on presynaptic axons (Venkatesan et al., 1996). Oke et al. (1997) examined the concentrations of NE and 5-HT in the human thalamus in postmortem studies. They found that NE was located particularly in the medial thalamus and ILN, with little in the pulvinar. 5-HT showed a similar distribution pattern in the anterior thalamus, but there were

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higher levels in the pulvinar that interestingly differed markedly between subjects. They found very little dopamine in the thalamus in line with previous animal studies. Papadopoulos and Parnavelas (1990) conducted a similar study in the rat lateral geniculate nucleus (LGN). They found that NE axons went mostly to the dorsal LGN, where they formed symmetrical synapses ( presumably inhibitory) on dendritic spines and shafts but not on somata. In contrast, 5-HT axons went mainly to the ventral LGN and intergeniculate leaflet, where they formed asymmetrical synapses ( presumably excitatory) on spines and dendritic shafts of both relay cells and Ins and symmetrical synapses on somata. The major excitatory input to the NE cells in the LC is glutamate, and the major inhibitory influence is GABA. Thirty-three percent of synapses on LC cells contain glu, and 66% contain GABA, and half the latter colocalize GABA and gly (Somogyi and Llewellyn-Smith, 2001). Bickford et al. (2000) found an interesting species difference in that the cat, but not the monkey, shows an NE projection from the LC to the LGN. The two species share other inputs to the LGN (i.e., ACh/nitric oxide [NO] from the PPN, GABA from the reticular nucleus of the thalamus and the pretectum, and ACh from the parabigeminal nucleus). Fung et al. (1994) describe another species difference in that some LC neurons in the rat, but not in the cat, contain enkephalin.

II. Functional Neuroanatomy

The experience of novelty results in the release of NE throughout the brain. In the prefrontal cortex and LC of conscious rats, this effect was mediated by 1 ARs. 2 ARs (as well as 5-HT[1A]Rs and GABA[A]Rs) had the opposite effect (Pudovkina et al., 2001). In the rat, thalamic ventrobasal nucleus, NE, released during behaviorally activated states, selectively enhances sensory transmission through lemniscal pathways by depolarizing the thalamocortical neurons and simultaneously inhibiting other sensory pathways (Castro-Alamancos, 2002). NE also enhances the signal-to-noise ratio by selectively inhibiting background neuronal activity. It does this by inhibiting EPSPs, particularly in the intracortically connected cortical layer Ib (Hasselmo et al., 1997). In finer detail, in cat visual cortex, NE modulates the spatiotemporal tuning of visual cortical neurons by enhancing their selectivity for stimulus speed and direction. In contrast, ACh simply increases and GABA decreases the amplitude of visually evoked responses. Neither alters the shape of the velocity tuning curve (McLean and Waterhouse, 1994). NE can stimulate or inhibit neurons or can exert a biphasic effect. NE release in the cortex is modulated presynaptically by a number of neuromodulators— inhibition by 2 ARs, MRs, and opioid and cannabinoid receptors—excitation

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by ARs, angiotensin, and bradykinin receptors (Schelb et al., 2001). These systems mature at different rates in the developing brain. There are two functionally distinct pools of synaptic vesicles at NE nerve terminals in the central nervous system (CNS). One is regulated by PKC and the other by dynamic changes in the cytoskeleton (Walaas, 1999). Activation of PKC potentiates stimulation-induced NE release (Schroeder et al., 1995). A. Excitatory Effects In rat brain thalamic slices, NE (and ACh) directly excites neurons in both the ILN and midline nuclei, as well as in the specific sensory neurons (Bayer et al., 2002). In rat SNpc slices, 1 ARs increase the spontaneous rate of firing of the SNpc cells (Berretta et al., 2000). In rat piriform cortex, NE released by stimulating LC neurons potentiates the response of the cortical neurons to odoriferous stimuli both in terms of spike count and temporal organization (Bouret and Sara, 2002). Similarly, stimulation of LC projections to the medial olfactory bulb potentiates the responses of the mitral cells to weak incoming stimuli from the nasal epithelium ( Jiang et al., 1996). The authors suggest that NE release improves the detection of relatively weak odors. In rat bed nucleus of the stria terminalis, NE fibers from the median forebrain bundle maintain an ongoing tonic activity in these neurons. Adequate levels of NE are maintained by modulating NE reuptake and by modulating NE release by way of presynaptic 2 ARs (Forray et al., 1997). B. Inhibitory Effects NE (and 5-HT) can inhibit neurons by two mechanisms (1) by increasing a Kþ conductance leading to hyperpolarization and (2) by activating GABA or Gly interneurons that evoke IPSPs in target neurons (Grudt et al., 1995). Infusion of 2 AR agonists into the LC in conscious rats greatly reduces NE levels in the PFC to which the LC projects. This indicates the presence and effectiveness of inhibitory 2A ARs on these LC cells. Infusions of N-methyl-d-aspartate (NMDA) into the LC increased levels of NE in the PFC by 240%. This provides evidence of excitatory NMDARs on these cells (Van Gaalen et al., 1997). The nucleus raphe magnus sends a predominantly inhibitory projection to the lateral reticular nucleus. Neurons in the latter respond to somatic stimuli on the dorsal body surface by inhibition. Glutamate applied iontophoretically to these cells also evokes an inhibitory response. In contrast, NE applied in this way to these cells inhibited only 9 of 21 neurons and had no effect on 11 of 21 neurons. However, in the latter case, NE blocked the inhibition induced


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by glutamate (Murphy and Behbehani, 1993). NE release in the pineal gland is subject to negative feedback control by presynaptic auto 2 ARs. This release exhibits a circadian rhythm (Drijfhout et al., 1996). C. Biphasic Effects Many neuromodulators can modulate the same system in one direction by acting on one set of receptors and can modulate the same system in the opposite direction by acting on a different set of receptors. For example, ascending projections from the LC to the thalamic parafascicular nucleus enhance nociception-activated neuronal activity acting on 1 ARs and can inhibit this activity by acting on 2 ARs (Zhang et al., 1998). In cat somatosensory cortex, Warren and Dykes (1996) applied NE by microiontophoresis in the vicinity of 31 neurons with receptive fields and 86 neurons without receptive fields while they were driven by glutamate. In 54% of cells of both types, NE was inhibitory, and in 36% it was excitatory. The former cells were located mainly in the middle layers of the cortex and the latter in the upper and lower layers. The authors suggest that the inhibiting effects of NE serve to increase the signalto-noise ratio in cortical processing, whereas its excitatory effects result in increased exchange of information with other cortical areas. In many cases, the effects lasted more than 5 minutes. The effect depended on 1 ARs. A similar biphasic effect was reported in the superficial layer of hamster superior colliculus. Here NE hyperpolarized 44% of cells tested. It did this by activating 2 ARs. NE depolarized 8% of cells tested by means of 1 ARs (Tan et al., 1999). In guinea-pig lateral septal slices, bath application of NE depolarized some septal neurons by means of 1 ARs and hyperpolarized others by means of 2 ARs. NE also induced rhythmic bursts by way of 1 ARs. The burst activity, as well as after polarization potentials, induced by NE is generated by a cationic conductance largely permeable to Naþ (Carette, 1999). In rat hypothalamic arcuate nucleus, NE excited neurons by means of 1 ARs and ARs. It also inhibited these neurons, possibly by means of an opioid-inhibitory neuron (Kang et al., 2000). The complexity of these systems is illustrated in an article by Parra et al. (1998). They found that inhibitory neurons in the HC differ widely in morphology and function. These authors described 16 different morphological phenotypes in the CA1 area and three different modes of discharge. Subtypes were either excited or inhibited by activation of NE, MR, 5-HT, and mGlu receptors. Most cells responded to two or three agonists, and 25 different response combinations were detected. However, the subsets defined by morphology, physiology, and pharmacology did not coincide, indicating that the hippocampal interneurons cannot be classified into a few well-defined groups.

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III. Electrical Activity

A. Potassium Channels 1. Postsynaptic In most neurons in the rat preoptic nucleus (PON), NE stimulated 2 ARs to induce membrane hyperpolarization associated with opening Kþ channels and an increase in membrane conductance. In some cases, NE depolarized PON neurons instead by means of 1 ARs (Bai and Renaud, 1998). A similar result was obtained in rat lateral septum slices (Liu and Alreja, 1998), medial pontine RF neurons where NE is ‘‘an excitatory neuromodulator’’ (Stevens et al., 1994), and rat hypoglossal motoneurons (Parkis et al., 1995). In the latter case, the authors implicated a rise in an inward Naþ current and a fall in a resting Kþ current. In rat HC CA1 slices, NE increased the frequency of IPSPs in pyramidal neurons by depolarizing surrounding INs. This was mediated by 1 ARs on the INs decreasing a Kþ conductance (Bergles et al., 1996). In mouse entorhinal cortex, stimulation of 2 ARs hyperpolarizes most neurons in layer II by promoting a Kþ conductance. This strongly inhibits glu transmission. In contrast, NE had no effect on IPSPs in this study (Pralong and Magistretti, 1995). NE also inhibits the Ca2þ-activated Kþ current IAHP that mediates slow afterhyperpolarizations and spike frequency adaption in hippocampal and neocortical neurons. This effect is mediated by 1 ARs, cAMP, and PKA (Pedarzani and Storm, 1996). Lancaster et al. (2001) found that short bursts of EPSPs can activate a Ca2þdependent Kþ current resembling I(sAHP) by activating VOCCs. The activation of this current reduces the postsynaptic response to high-frequency synaptic input. Application of NE on rat HC CA1 pyramidal cells reduced the resting slow afterhyperpolarization (sAHP) conductance (Lancaster et al., 2001). Wagner et al. (2001) also found in guinea pig POA that NE inhibited an apamin-sensitive small conductance Ca2þ-activated Kþ channel that underlies at least part of the I(AHP) current in this location. In rat SNpc, both NE and DA inhibit the hyperpolarization-activated cationic (Ih) current and induce an inwardly rectifying Kþ (I(Kir)) current. PLA, PKC, and phosphatases 1 and 2A are not involved in this effect (Cathala and Paupardin-Tritsch, 1999). Hypoglossal motoneurons possess TASK-1, which is a two-pore domain Kþ channel that provides a prominent leak Kþ current. This channel is inhibited by NE and by 5-HT. This inhibition of a hyperpolarizing current induces a slow excitation of the neuron (Talley et al., 2000). 2. Presynaptic NE cells in the A5 noradrenergic area have 2 ARs, activation of which increases an inwardly rectifying Kþ current but has no effect on Naþ currents (Huangfu and Guyenet, 1997).


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B. Calcium Channels In many areas of the brain, NE suppresses calcium currents. 1. Presynaptic In Xenopus olfactory bulb, presynaptic 2 ARs inhibit Ca2þ currents in mitral cells but not in granule cells (Czesnik et al., 2001). In rat HT synaptosomes, presynaptic ARs inhibit NE release by reducing Ca2þ influx through VOCCs (Martire et al., 1995). In rat cortical synaptosomes, Kþ-evoked NE release is mediated by activation of P/Q- and N-type Ca2þ channels (Grassi et al., 1999) or L-type Ca2þ channels (Sabria et al., 1995). In rat HC, NE release is mediated by N-type Ca2þ channels (Sabria et al., 1995). Interestingly, autoantibodies present in the sera of patients with ALS can interact with these channels, leading to an increase of potentially neurotoxic Ca2þ influx and to excess NE release (Grassi et al., 1999). In rat brain synaptosomes, ischemia reverses the Naþ/Ca2þ exchanger, leading to increased i.c. Ca2þ levels and increased NE release (Du et al., 1997). 2. Postsynaptic In rat nucleus of the solitary tract, NE inhibits N- and P-type high-voltage activated Ca2þ channels by means of a PTX-sensitive G-protein (Ishibashi and Akaike, 1995). Bulbospinal neurons in the rat rostroventral medulla exhibit two types of calcium current. One is activated by high voltages and the other by low voltages. NE inhibits the former by means of 2 ARs but not the latter. The NE-sensitive currents were mainly of the N- and P/Q-types (Li et al., 1998). Kolaj and Renaud (2001) found that stimulation of postsynaptic 2 ARs in the rat PON suppressed mainly N-type Ca2þ channels by means of a PTX-sensitive G-protein, as well as some L-and P/Q-type Ca2þ channels. NE can also evoke the release of Ca2þ from internal stores. In rat pineal cells, NE induces an increase in i.c. Ca2þ levels in two stages: (1) by a rapid release of Ca2þ from internal stores with a single or repeated spikes of Ca2þ concentration and (2) by a later and more prolonged increased influx of Ca2þ leading to a plateau. In contrast, ACh promotes a single abrupt rise in i.c. Ca2þ levels followed by an exponential fall. Furthermore, the elevations in i.c. Ca2þ levels induced by NE and ACh derive from different i.c. stores (Marin et al., 1996). 3. Miscellaneous Items In cultured astrocytes, NE (and glu) induce i.c. transients, oscillations, and waves. Furthermore, NE, by means of 1 ARs, mediates neuron-astrocyte communication (Duffy and Mac Vicar, 1995). NE induces Ca2þ oscillations by a G-protein–dependent PI pathway in which Ca2þ is released from inositol 1,4,5 triphosphate-sensitive stores (Tse and Tse, 1998). This mechanism underlies the action of NE in releasing ACTH (Tse and Tse, 1998). In rat

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TABLE 1 Effect of NE, GABA, and 5HT on Three Parameters of the Rhythmic Slow Activity Induced by ACh

Interburst interval Burst duration Total power

NE

GABA

5HT

þ þ þ

þ 0 –

þ þ –

pituitary intermediate lobe cells, NE (by means of -adrenergic receptors) induces first a slight increase, and then a marked decrease, in i.c. Ca2þ levels, leading to the secretion of -endorphin (Nemethy et al., 1998). In rat HC slices, NE modulates the rhythmic slow activity induced by ACh by increasing interburst intervals, burst duration, and total power (Table I) (Boguszewicz et al., 1996).

IV. Interactions with Other Transmitters

A. Norepinephrine and Glutamate 1. NE ! Glu a. Presynaptic. NE regulates glutamate (glu) release in many regions either up or down. For example, in rat auditory brain (medial nucleus of the trapezoid body), stimulation of 2 ARs inhibits glu release by downregulating Ca2þ currents (Leao and Von Gersdorff, 2002). In rat bed nucleus of the stria terminalis, NE maintains an inhibitory tone over the information flow through the nucleus carried by glu neurotransmission. It does this by inhibiting Kþ-induced glu release by means of 2 ARs and by inhibiting its own release by means of 2 ARs (Forray et al., 1999). In rat HC synapses, NE reduces excitatory (glu) but not inhibitory currents by means of inhibitory 2 ARs on glu terminals. This effect involves inhibition of VOCCs and not by means of modulating spontaneous vesicle exocytosis or voltage-activated Kþ channels (Boehm, 1999). Blockade of 2 ARs potentiates bursting responses of LC neurons to locally applied glutamate (Simson, 2001). In rat spinal cord, 2A/D subtypes of the AR inhibit capsaicin-induced glu release, thus inhibiting pain (Li and Eisenach, 2001). Also in rat spinal cord, 2A adrenergic receptors on axon terminals located on sympathetic preganglionic neurons inhibit glu release (Miyazaki et al., 1998). Activation of PKC by NE inhibits group III mGluRs and increases excitatory


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synaptic input in oxytocin and vasopressin neurons in the paraventricular nucleus of the hypothalamus (Gordon and Bains, 2003). This results in an increase in mEPSC frequency in response to a subsequent exposure to NE. The authors suggest that, by removing feedback inhibition, this mechanism effectively primes the synapses such that subsequent activation is more efficacious. ‘‘The novel form of synaptic rescaling afforded by this cross-talk between distinct metabotropic receptors provides a means by which ascending catecholamine inputs can facilitate the control of homeostasis by hypothalamic networks’’ (Gordon and Bains, 2003). NE can also potentiate glu release, as in the case of 1 ARs (and 5-HT(2 ARs) on glu terminals located on apical dendrites of layer V pyramidal cells in rat medial PFC (Marek and Aghajanian, 1999). In rat LC, glutamate applied iontophoretically to distal dendrites potently and directly activated LC neurons. This effect was further potentiated by stimulation of 2 ARs (Ivanov and Aston-Jones, 1995). b. Postsynaptic. NE and glu also interact at postsynaptic sites. In rat embryonic visual cortex, Yang et al. (1996) measured i.c. Ca2þ levels. The addition of NE alone had no effect, and the addition of glu had only a small effect. However, both together caused a marked rise (up to 8) in i.c. Ca2þ levels. This effect was mediated by NMDARs and ARs. The authors suggested that this result indicates that ARs and NMDARs colocalize and interact at postsynaptic sites on cortical neurons. In sensory cortex neurons, NE (by means of 1 ARs) potentiates glu transmission and (by means of ARs) inhibits glu transmission both at postsynaptic sites (Devilbiss and Waterhouse, 2000). c. Further Biphasic Effects. In rat basolateral amygdala slices, NE inhibited both fast and slow EPSPs by means of 2 ARs and to a lesser extent increased both types of EPSP by means of ARs (Ferry et al., 1997). In the parvocellular region of rat paraventricular nucleus of the hypothalamus, NE caused a marked increase in the frequency of EPSPs in 36% of cells examined (by means of 1 ARs acting to increase glu release) and a direct hyperpolarizing inhibitory response (by means of ARs) in 14% of cells examined. The excitatory response was indirect, being mediated by NE-sensitive glutamatergic interneurons, but the inhibitory effect was direct. This system modulates the release of pituitary hormones (Daftary et al., 2000). NE can also affect glu synthesis. In mouse astrocytes, NE promotes glutamine uptake, glu synthesis, and CO2 production, thus increasing the energy supply to astrocytes (Huang and Hertz, 1995). 2. Glu ! NE In rat supraoptic nucleus (SON), activation of presynaptic NMDARs inhibits a NE release that depends on mobilization of intracellular Ca2þ stores (Liu et al., 2001). In contrast, activation of presynaptic NMDARs in the bed nucleus of the

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stria terminalis promotes NE release (Forray et al., 1995). In rat HC synaptosomes, activation of presynaptic glu receptors also promotes Ca2þ-dependent release of NE (and DA) (Malva et al., 1994). In rat HC slices, NMDA promoted NE release from the dentate gyrus but not from the CA1–CA3 area (Andres et al., 1993). Von Kugelgen and Starke (1995) added to the complexity by their finding that, in rabbit cortex, activation of kainate (KA) glu receptors (but not AMPA, NMDA, or mGlu receptors) inhibits NE release indirectly by means of the release of adenosine, which stimulates presynaptic inhibitory adenosine A1 receptors. In rat striatum, Ohta et al. (1994) report that activation of both NMDA and AMPA/KA receptors stimulated NE, DA, and 5-HT release. Each type of gluR exerted a differential effect on the release of these three neuromodulators. Pile (1996) report that rat brain contains mGluRs, activation of which can either inhibit or enhance NE-stimulated cAMP accumulation, depending on circumstances. Enhancement was mediated by means of adenosine and inhibition by means of receptors sensitive to quisqualate. In rat hypothalamic slices, glu stimulates NE release. This release was ‘‘dramatically’’ reduced by GABA(B)R activation (Navarro et al., 1995). The rat PFC exerts a potent excitatory input (presumably glutamatergic) onto LC neurons and provides a resting tonic excitatory influence ( Jodo et al., 1998). B. Norepinephrine and GABA 1. NE ! GABA a. Presynaptic. NE has varied actions on GABA release, depending on receptor type. Local infusions of NE inhibit GABA release in the rat SON by means of presynaptic ,2 ARs (Wang et al., 1998) but promote GABA release in human, rat, and guinea-pig cortical slices and synaptosomes by means of 1 ARs (Ferraro et al., 1993) and also promote GABA release in kitten visual cortex by means of 1 ARs (Shirokawa and Ogawa, 1994). In rat LC, Tamiya et al. (1994) found that a single NE neuron simultaneously receives synaptic inputs from GABA and substance P afferents. These NE neurons made feedback symmetrical synaptic contacts with the GABA axon terminals and asymmetrical synaptic contacts with the substance P terminals. In midbrain DA neurons, activation of presynaptic NE ARs inhibits the IPSPs evoked by GABA. 5-HT and DA also had this effect (Federici et al., 2002). b. Postsynaptic. NE acting postsynaptically generally promotes GABA transmission. In rat cerebellum, NE induces a long-term facilitation of GABAergic transmission. This is effected by excitatory 2 ARs on the GABAergic basket cells that in turn evoke IPSPs on Purkinje cells (Saitow et al., 2000). Mitoma and Konishi (1999) found that in rat cerebellar slices, NE (and 5-HT ) evokes longterm potentiation of GABAergic transmission by means of a cAMP-protein


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kinase-dependent mechanism. Sessler et al. (1995) confirm that in rat somatosensory cortex layer V pyramidal neurons, NE potentiates GABAergic inhibition by means of a AR cAMP-dependent mechanism. Similarly, in medial septal/ diagonal band (MS/DB) neurons, NE excites GABAergic neurons by means of 1 ARs (Alreja and Liu, 1996). In rat spinal cord substantia gelatinosa, NE excites GABAergic INs by way of postsynaptic 1 ARs on somata and dendrites (Baba et al., 2000). In the rat basolateral nucleus of the amygdala, NE excites GABAergic INs by way of asymmetrical synapses located on medium and small dendrites (Li et al,. 2002). c. Bimodal. In the guinea-pig lateral septum (LS), NE axons from the LC act on LS neurons in two ways—a direct excitatory effect by means of both 1 and 2 receptors and an indirect inhibitory effect mediated by stimulating inhibitory GABAergic INs (Carette et al., 2001). A different bimodal arrangement is found in the rat hypothalamic PVN (Han et al., 2002). In this case, NE exerts a dual control of the tonic GABAergic input from local INs to most type II PVN neurons. Stimulation of excitatory postsynaptic 1 ARs on the somata and dendrites of type II PVN cells increases the frequency of IPSCs. In addition, stimulation of inhibitory presynaptic 2 ARs on GABAergic terminals decreases the frequency of IPSPs by preventing GABA release. NE also increases the frequency of EPSPs in this location (Han et al., 2002). In rat substantia nigra, NE both activates some 50% of DA neurons by inducing EPSCs by means of 2 ARs and also stimulates presynaptic 2 ARs to promote GABA release that leads to an increase of IPSC frequency (Cathala et al., 2002). In rat red nucleus, NE (by means of 2 ARs) increases the response of rubral neurons to GABA and also (by means of ARs) decreases this response (Ciranna et al., 2000). d. Miscellaneous. The postsynaptic cascade by which NE potentiates the amplitude of GABA(A) responses in rat cerebellar Purkinje calls runs ! G-protein ! adenylate cyclase ! cAMP ! phosphorylation of the GABA(A) receptor by cAMP-dependent cyclase (Cheun and Yeh, 1996). NE modulates the synthesis of GABA in central limbic stress circuits. A decrease in GABA synthesis is related to a decreased activation of stress– excitatory pathways and to the enhanced activation of stress inhibitory pathways (Herman et al., 2003). This suggests that NE carries the signal ‘‘stress’’ to activate GABAergic mechanisms in the limbic system. The response of GABAergic INs in rat frontal cortex to NE depends on their type (Kawaguchi and Shindou, 1998). Common multipolar, chandelier, and neurogliaform cells responded with depolarization without spike firing. Somatostatin-positive Martinotti and wide arbor cells were also depolarized but produced spikes in addition. CCK-positive cells (including large basket cells) were affected in various ways—depolarization, hyperpolarization, or both sequentially.

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2. GABA ! NE a. Excitation. In rat brain slices, activation of GABA(A) receptors augments electrically stimulated NE release by means of disinhibition of INs (Fiber and Etgen, 1998, 2001). This effect is inhibited by opioids and by adenosine (Fiber and Etgen, 2001). In rat LC, single NE neurons can receive afferent fibers from both GABA and SP neurons. These NE neurons then make symmetrical (inhibitory) synaptic contacts with the incoming GABA terminals and asymmetrical (excitatory) contacts with the incoming SP axon terminals (Tamiga et al., 1994). In rat HC synaptosomes, Fassio et al. (1999) report that GABA potentiates NE release by two mechanisms (1) activation of GABA(A)Rs located presynaptically on NE terminals leading to NE release by exocytosis and (2) by activating GABA transporters. Process (1) requires N- and P/Q-type Ca2þ channels, and process (2) needs N-type Ca2þ channels only. b. Inhibition. NE neurons in the nucleus of the solitary tract (NTS) project to the subfornical organ. This projection is tonically inhibited by local GABA neurons in the NTS by means of GABA(A) receptors (Tanaka et al., 2002). In rat red nucleus, GABA diminishes the response of rubral neurons to NE (Ciranna et al., 2000). c. Bimodal. In rat brain synaptosomes, GABA enhances basal NE release but inhibits Kþ-evoked NE release. The former effect is associated with an increase in Ca2þ entry and is coupled with a Cl–/Ca2þ symporter system (Cerrito et al., 1998). 3. NE ! ACh a. Postsynaptic. Postsynaptically, NE inhibits some 90% of rat brainstem cholinergic neurons directly by activating the inwardly rectifying Kþ current IKG (Williams and Reiner, 1993). In rat brain slices, NE directly inhibits cholinergic neurons in the LDTN but excites noncholinergic local circuit INs by two distinct mechanisms—one direct and the other long-lasting and indirect. The latter was mediated by glu neurons (Kohlmeier and Reiner, 1999). In contrast, NE depolarizes striatal cholinergic interneurons by means of 1-adrenoceptor activation through a cAMP-dependent, but PKA-independent, mechanism (Pisani et al., 2003). 4. NE ! 5-HT Dorsal raphe 5-HT neurons are activated by three arousal-related mechanisms including NE (as well as histamine and orexin/hypocretin). This effect is probably mediated by opening a mixed cation channel (Naþ, Kþ) (Brown et al., 2002*). Stimulation of postsynaptic 2 ARs on raphe 5-HT neurons leads to the inhibition of (1) N- and P/Q-type Ca2þ channels but has no action on (2) G-protein-coupled inwardly rectifying Kþ channels. In contrast, 5-HT potentiates (2) as well as inhibiting (1) (Li and Bayliss, 1998). 2 ARs, in rat HC (Numazawa et al., 1995) and in rat HC and cerebral cortex (Scheibner et al., 2001) inhibit Kþ-evoked 5-HT release by means of a G-protein.


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In contrast, in rat pinealocytes in culture, NE triggers Ca2þ-dependent exocytotic release of 5-HT as the first step in melatonin synthesis (Yamada et al., 2002). In rat basolateral amygdala, stimulation of ARs promotes synaptic transmission by means of cAMP and PKC. This effect is inhibited by stimulation of 5-HT (1A)Rs by action at a site downstream from cAMP production ( Wang et al., 1999). In rabbit raphe, there are two mechanisms for upregulation by NE of the firing rate of 5-HT neurons: (1) direct excitation by means of 1 ARs on the 5-HT neuron and (2) by disinhibition by means of inhibitory 2 ARs on the inhibitory collateral feedback loop from the 5-HT axon onto the dendrites of the same 5-HT cell (Kalsner and Abdali, 2001).

V. Postsynaptic Cascades

A. cAMP The cAMP pathway plays a role in many of the effects of NE. For example, in rat hippocampal CA1, pyramidal neuron stimulation of ARs activates cAMP-dependant kinase and so enhances EPSCs (Gereau and Conn, 1994). In contrast, in rat cerebellar stellate cells, NE enhances IPSCs by means of ARs (Llano and Gerschenfeld, 1993). The signaling pathway for this effect runs ! activation of adenylate cyclase ! increased cAMP levels ! activation of protein kinase A ! modulation of the vesicle release mechanism rather than by modulating i.c. Ca2þ levels (Kondo and Marty, 1997). Elevation of cAMP levels also reduces the afterhyperpolarization after single-action potentials in NE neurons. This effect is not dependent on PKA (Shiekhatter and Aston-Jones, 1994). NE is the most potent stimulator known of melatonin synthesis in the pineal gland. Maronde et al. (1997) showed that this is mediated by the cascade: ARs ! cAMP ! cAMP-dependent kinase II ! phosphorylation of CREB. In bovine pineal gland, NE application induces cAMP production that serves to protect arylalkylamine N-acetyltransferase (the key enzyme regulating melatonin production) from proteolytic destruction (Schomerus et al., 2000). In rat pinealocytes, the production of cAMP by NE is modulated by pretreatment with fatty acids— augmentation by eicosapentaenoic acid and inhibition by docosahexaenoic acid (Delton-Vandenbroucke et al., 1996). NE activates melatonin secretion from rat pinealocytes. These cells also release glu, which activates autoreceptors in a paracrine manner. This serves to inhibit NE-stimulated melatonin release (Yamada et al., 1996). NE also interacts with glu and GABA mechanisms. In rat cerebral cortical slices, different types of mGluRs either potentiate or inhibit the NE-stimulated accumulation of cAMP. The potentiation is mediated by mGluRs blocked by

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L-AP3 and glutamate; the inhibition is mediated by receptors sensitive to quisqualate (Pilc et al., 1996). In hippocampal neurons, Raman et al. (1996) showed that stimulation of ARs during excitatory synaptic transmission can increase charge transfer and Ca2þ influx through NMDARs. This is mediated by promoting recovery of NMDARs from the calcineurin-mediated dephosphorylation induced by synaptic activity. In HC dentate cells, NE-activated cAMP-dependent protein kinase enhances GABA(A) currents (Kapur and Macdonald, 1996). In rat cerebellar Purkinje cells, stimulation of ARs also enhances GABA(A)R function by means of the chain G-protein ! cAMP ! and increased phosphorylation of the GABA(A)R by cAMP-dependent protein kinase (Cheun and Yeh, 1996). In rat somatosensory cortical layer V pyramidal cells, the enhancement of GABAergic responses by stimulation of ARs is masked to some extent by inhibition of these responses induced by AR stimulation (Sessler et al., 1995). In rat HC CA1 cells, activation of ARs starts a signaling cascade that runs ! G-alpha ! adenylate cyclase ! increased cAMP levels ! activation of cAMP-dependent protein kinase ! phosphorylation and activation of the Naþ-Hþ exchange and so to an increase of i.c. pH. (Smith et al., 1998). In the thalamus, the cascade starting with NE and resulting in LHRH release runs ! NE ! NOS activation ! NO release ! PGE2 synthesis ! activation of adenylate cyclase and cAMP formation ! exocytosis of secretory granules for LHRH (Rettori et al., 1992). In rat cerebral cortex, both NE and E activate Ars, which leads to the secretion of cAMP from astrocytes and its hydrolysis in extracellular space to produce the neuromodulator adenosine (Rosenberg and Li, 1995). In rat olfactory bulb, Yuan et al. (2003) show that three cascades converge on mitral cells to modulate learning. These are (1) the excitatory pathway involving raising i.c. Ca2þ levels from the nasal epithelium activated by odor (2) the cascade NE-cAMP involving 1 ARs, and (3) a cascade involving 5-HT(2A)Rs. The end result of this convergence is to modulate the phosphorylation of CREB. The NE-cAMP cascade may also be involved in Alzheimer’s disease (AD). Neurotransmitters, which activate phosphatidylinositol hydrolysis and PKC, stimulate APP processing and generate soluble, nontoxic nonamyloidogenic peptides. Elevations in cAMP levels, produced inter alia by NE, oppose this effect and lead to the accumulation of toxic amyloidogenic A peptides (Lee et al., 1997). B. MAPK/ERK In rat pinealocytes, stimulation of both and ARs activates the MAPK signaling pathway. This involves two separate but integrated pathways—one by way of PKA and the other by way of PKG—that have opposite effects on MAPK phosphorylation (Ho and Chik, 2000).


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The mood-stabilizing agents lithium and valproic acid activate ERK. This regulates AP-1 function and so controls the expression of several genes, including those for growth-cone–associated protein 43 and Bcl-2. This activity promotes neurite growth and cell survival and also enhances NE uptake and release (Yuan et al., 2001). C. Calcineurin In rat brain, NE stimulates 1 Ars, causing a rise in i.c. Ca2þ levels. This initiates the path running ! activation of the calmodulin-dependent phosphatase calcineurin ! activation by dephosphorylation of NAþ-Kþ ATPase (Mallick et al., 2000). D. Phosphatidylinositol Many of the effects of NE are mediated by the inositol phosphate signaling cascade that includes a G-protein and PLC. PLC hydrolyzes phosphatidyl inositol 4,5-biphosphate (PI) to the second messengers inositol phosphate (IP) and 1,2diacylglycerol (DAG). DAG then releases Ca2þ from internal stores. In rat thalamic slices, NE stimulates 1 and 2 Ars, resulting in increased i.c. IP levels (Trejo et al., 1996). In rat striatum, stimulation of 1 ARs also evokes increased Ca2þ influx, activation of PLC, and raised i.c. (IP) levels (Arias-Montano et al., 1996). Marin et al. (1993) describe a complex PI-based signaling system in mouse striatal astrocytes. NE stimulates 2 ARs linked to PLC. Activation of PLC raises i.c. IP levels by means of a PTX-sensitive G-protein and also increases i.c. Ca2þ levels by means of DAG. The raised level of i.c Ca2þ activates PLA, which hydrolyzes membrane lipids to release arachidonic acid (AA). The AA released then inhibits glu uptake into the astrocytes that, in turn, leads to the stimulation of mGluRs coupled to PLC. A variation on this system is reported by Kreda et al. (2001) in immortalized HC neurons. Here stimulation of 1 ARs activates PLC to produce IP and DAG. DAG releases Ca2þ from internal stores, and this activates PLA2, leading to AA release and secretion of luteinizing hormone-releasing hormone (LHRH). However, in LA-N-2 cells, NE induces raised IP levels by a different mechanism. This includes a G-protein and PLC but not Ca2þ (Sorrentino et al., 1996). E. Prostaglandins Another cascade influenced by NE is mediated by the sequence G-protein ! phospholipase A2 ! cyclooxygenase (COX) and a prostaglandin. Prostaglandin receptors of various types occur on NE neurons. For example, PGE3 receptors

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occur on most NE (as well as 5-HT and adrenergic, but not DA) neurons (Nakamura et al., 2001). In rat HC mossy fiber nerve endings, PLA2 mediates the synthesis of PGF2. This PLA2 is activated by NE, 5-HT, and DA (but not by ACh, glu, GABA, histamine, or purines) (Separovic and Dorman, 1993). In rat HT, IL-1 activates COX to synthesize PGD2, which then activates NE neurons (Terao et al., 1995). Likewise, in rat HT PVN, the release of NE by IL-1 is mediated by prostaglandin synthesis (Tsumori et al., 1998). Stimulation of the PLA2–Cox cascade leads to activation of the sympathoadrenomedullary outflow, in which process brain thromboxane A(2) is involved (Yokotani et al., 2000). COX (cyclooxygenase) is a potent source of ROS. F. Nitric Oxide Nitric oxide (NO) is involved in a vast number of activities in the brain. These include effects initiated by NE. 1. Presynaptic NO evokes the presynaptic release of NE under a variety of conditions. In synaptosomes, NO evokes NE release both directly from terminals and indirectly by releasing glutamate (Lonart and Johnson, 1995). In rat brain slices, NO enhances NMDA-stimulated NE release by inhibiting cellular respiration and perhaps ultimately by altering calcium homeostasis (Stout and Woodward, 1995). In rat cortical synaptosomes, NO evokes NE release by means of activation of guanylate cyclase under both basal and Kþ-evoked depolarization conditions (Martire et al., 1998). In rat HC, NO evokes NE release in the presence of thiol compounds, such as l-cysteine, in a calmodulin-dependent but cGMPindependent manner (Satoh et al., 1996). On the other hand, the application of NMDA to cholinergic neurons in the pons resulted in an increase of NO levels and a decrease in NE release (Kodama and Honda, 1999). In rat HT, angiotensin inhibits NE release by means of an NO mechanism acting on guanylcyclase, but in the peripheral nervous system, angiotensin promotes NE release acting presynaptically (Gironacci et al., 2000). 2. Postsynaptic Postsynaptically, NO is involved in NE mechanisms in a number of ways. For example, in the rat, intracerebroventricular injection of NO donors activates COX and promotes PG synthesis in the NE neurons that innervate the PVN, resulting in an increased level of blood corticosterone (Okada et al., 2002). A similar signaling cascade is involved in the NE-stimulated release of luteinizing hormone releasing-hormone (LHRH) described previously. This cascade runs NE ! NOS ! COX ! PGE2 ! activation of adenylate cyclase !


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cAMP! exocytosis of LHRH secretory granules (Rettori et al., 1992). Seilicovich et al. (1995) report a close variation of this cascade in rat HT that uses cGMP instead of cAMP. In rat pineal gland, NE acting on ARs inhibits the firing rate of a population of neurons by means of NO that seems to act either directly on the electrical discharge of the cell or indirectly by means of cGMP (Schenda and Vollrath, 1998). Vacher et al. (2003) present evidence to show that the effect of NO on magnocellular oxytocinergic neurons in the PVN and SON may be exerted by modulating their main afferents, which includes NE axons, rather than by a direct action on the magnocellular neurons themselves. In rat cerebral astrocytes, stimulation of ARs induces the synthesis of NO. In 75%, the receptors concerned are , and in 25% they are (Agullo and Garcia, 1992). 3. More on cGMP Most neurotransmitters that raise cGMP levels do so by means of NOS. However, Morton and Bredt (1998) showed that NE also raises cGMP levels in NOS knockout mice in both granule cells and astrocytes in the cerebellum. This suggests that there must be a non-NOS cGMP pathway in the nervous system. G. Reactive Oxygen Species There is abundant evidence on the role of redox factors at the glutamate synapse (Smythies, 2002) but very little such information at the NE synapse. In fact, my Pub MED search recorded only two hits. Langeveld et al. (1995) report that H202 induces a nonexocytotic release of catecholamines (DA > NE) by interfering with vesicular uptake and storage. Kimelberg and Goderie (1993) found that in neonatal rat cerebral cortex, the antioxidant ascorbate increased Naþ-dependent NE uptake but inhibited Naþ-independent NE uptake. They also found a species difference in that mice, unlike rats, do not possess a Naþ-dependent NE uptake. However, several articles report on redox-related NE neurotoxicity and neuroprotection. Catecholamines, in general, can protect against or induce oxidative stress, depending on circumstances. The oxidative stress can result in apoptosis. Zilkha-Falb et al. (1997) tested catecholamines and serotonin on chick embryo sympathetic neurons and found them all to induce apoptosis with the rank order DA > NE > E > 5-HT. The authors present evidence to show that this effect is mediated by toxic o-quinone metabolites. Lymphocytes can also produce catecholamines that can act as autocrine and/or paracrine regulators of apoptosis ( Josephsson et al., 1996). In mouse cortical neuron cultures, catecholamines at high doses (>100 M ) are neurotoxic, producing apoptosis, whereas at low doses (1–30 M ), they protect against ROS-induced neurotoxicity by acting as antioxidants (Noh et al., 1999). In PC12 cells, catecholamines (DA > NE) are

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neuroprotective by virtue of their ability to scavenge ROS (Sofic et al., 2001). In rat cortical neurons, catecholamines protect against cell death partly by reducing the oxidative stress induced by the inhibitory action of glu on the cytosineglutamate transporter. Inhibition of this transporter results in a decline in i.c. levels of glutathione, which is the principal i.c. antioxidant in the brain (Ishige et al., 2001). An interesting finding (Troadec et al., 2002) was that NE protects DA cells (function and survival) by its own antioxidant function. This involves the autooxidation of NE to o-quinone metabolites. Khorchid et al. (2002) found that in cultured oligodendrocytes, NE (and DA) induced oxidative stress as determined by increased ROS production and reduced i.c. glutathione levels. This resulted in apoptosis. Stimulation of ARs in heart myocytes also can lead to apoptosis. The mechanism involves the sequential production of ROS, activation of JNK, and the mitochondrial death pathway (Remondino et al., 2003). Activation of 2 ARs on astrocytes protects against glutamate-induced cell death ( Junker et al., 2002). In rat glioma cell lines, NE, by means of ARs, protects against vitamin D3-induced apoptosis (Canova et al., 1997). NE oxidation in the presence of l-cysteine leads to the synthesis of dihydrobenzothiazide. This compound is an inhibitor of mitochondrial complex 1 and of other mitochondrial enzymes such as CoQ(1) reductase, pyruvate dehydrogenase, and -ketoglutarate dehydrogenase. This process may be related to the degeneration of neuromelanin-pigmented neurons in the LC in Parkinson’s disease (Xin et al., 2000). H. Endocytosis In contrast to other receptors, there is very little information on the role of endocytosis at NE receptors. In mouse T3 cells, 1B ARs are endocytosed by a mechanism in which PLC and PKC play a role but changes in i.c. Ca2þ do not (Awaji et al., 1998). Using 1B-adrenoceptor––inducible DDT1MF-2 cells for the sorting process and CHO cells stably expressing B-adrenoceptors for the agonist-promoted endocytosis, this same group (Hirasawa et al., 1998) showed that the endocytosis of 1B ARs depends on actin polymerization, whereas the return of these receptors to the plasma membrane requires intact microtubule function. I. Synaptic Plasticity In human neuroblastoma cells in culture, the addition of NE to the medium increased the number of neurites, as well as raised GAP-43 levels (a neural growth cone marker). This treatment also increased the expression of 44 genes,


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including NCAM L1 and laminin (Laifenfeld et al., 2002). Stimulation of the rat locus ceruleus results in NE release in the dentate gyrus. This produces potentiation of granule cells of the dentate gyrus both on a short-term (3 hours) and longterm (24 hours) scale. However, it produced a potentiation of the field EPSP in the dentate gyrus only at the long-term period. This potentiation was dependent on new protein synthesis ( Walling and Harley, 2004). The authors suggest that NE may facilitate long-term and not short-term memory. In an electron microscopic study, Matsukawa et al. (2003) showed that depletion of NE and/or 5-HT in the rat visual cortex leads to loss of synapses (29–55%), particularly in neonatal animals (1–2 weeks pp). J. Genes Activation of NE receptors triggers many cascades that lead to changes in gene expression as detailed elsewhere in this book under several topic headings. Here are some other such effects. NE regulates melatonin production and many other aspects of pineal function through actions involving cAMP and the stimulation of cAMP response element-binding protein (CREB) (Roseboom and Klein, 1995). Local infusion of NE increases c-fos expression in the mouse amygdala (Stone et al., 1997). Activation of the DM thalamic nucleus induces c-fos expression in its projection area in rat PFC in freely moving rats. This effect is strongly potentiated in catecholamine-depleted animals (Bubser et al., 1997) The authors suggest that this result indicates that catecholamines modulate the degree to which thalamic activity can activate the PFC in awake animals. NE can inhibit the expression of inflammatory genes in glial cells. Moreover, stimulation of ARs in astrocytes increases the expression of inhibitory 1k protein in rat frontal cortex (Garrilyuk et al., 2002). Opiates increase c-fos expression and NE turnover in the PVA (Laorden et al., 2000).

VI. Tropins and Cytokines

A. Tropins There are several reported interactions between the NE system and tropins. Stimulation of 2 ARs in rat HC neurons raises constitutive levels of TNF mRNA and protein (Renauld and Spengler, 2002). NE neurons, which project from the rat lateral septum through the medial forebrain bundle, contain BDNF that is anterogradely transported to the terminal where it is co-released with NE.

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The BDNF then promotes the expression of calbindin in the target cell, as well as promoting the arborization of the calbindin-containing neurons (Fawcett et al., 2000). Chronic opiate exposure leads to the upregulation of the cAMP signaling pathway and increased tyrosine hydroxylase expression, thus increasing NE synthesis. The presence of BDNF is essential for this effect (Akbarian et al., 2002). In goldfish pituitary cells, stimulation of 2 ARs inhibits the release of growth hormone by means of cAMP (Yunker et al., 2000). In rat brown adipose tissue, stimulation of ARs increases the expression of fibroblast growth factor (FGF ) 2 (Yamashita et al., 1995). B. Cytokines The i.p. injection of IL–1 in mice results in the appearance of the immediate early gene c-fos in several brain areas, particularly the HT PVN. This indicates that these areas were stimulated. This effect is substantially reduced by pretreatment with 6-OH DA, which depletes cerebral NE. NE innervation of the PVN is, therefore, likely to be involved in this effect (Swiergiel et al., 1996). IL-1 activates central sympathetic outflow, leading to raised NE plasma levels by means of a cascade in which PGs and NO are involved (Murakami et al., 1996; 2002). IL-1 infused into rats by microdialysis activates the glu system, leading to activation of COX and NOS, which results in increased NE release (Kamikawa et al., 1998). IL-1 promotes the release of NE and 5-HT by means of the COX-AA signaling pathway (Mefford and Heyes, 1990).

VII. Memory and Learning (LTP)

Experiments with TH knockout mice show that NE is essential for the consolidation process of long-term memory (Kobayashi, 2001). In rat visual cortex, NE-mediated rises in i.c. Ca2þ levels promote LTP at inhibitory and at excitatory synapses (Komatsu and Yoshimura, 2000). In rat HC slices, NE, by means of 1 ARs, promotes LTP (whereas ACh, NMDA, and mGlu agonists do not) (Izumi and Zorumski, 1999). Perfusion of HC slices with a AR agonist induces LTP of responses evoked by stimulation of the perforant path and LPD of responses evoked by stimulation of the lateral perforant path (Pelletier et al., 1994). Yang et al. (2002) showed that rats with 6-OH DA lesions in the HC CA1 area lose the ability to develop LTP but not LTD. Therefore, in this location NE may be important for LTP but not LTD. In rat visual cortical slices, neither paired pulse stimulation (PPS) nor stimulation of M1Rs, or 1 ARs, produced LTD by themselves, but PPS plus


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one of the others together did so. This effect requires NMDARs (Kirkwood et al., 1999). The authors conclude the NE controls the gain in homeostatic-dependent homosynaptic LTD in visual cortex. Some forms of olfactory learning depend on NE terminals acting on receptors in the olfactory bulb that modulate the dendrodendritic inhibition of mitral cells by granule cells by producing a depression-potentiation sequence in granule cell–mediated inhibition of mitral cells (Okutani et al., 1998). Medial frontal lesions in postnatal 7 day rats cause defects in spatial navigation task performance that recover over time. This recovery is associated with an increase in spine numbers per unit length of the dendrites of cortical pyramidal cells. Depletion of cerebral NE before the lesion prevents this recovery (Kolb et al., 1997). Locus ceruleus–initiated synaptic potentiation in rat dentate gyrus (by means of the perforant path) may selectively support long-term, rather than short-term, memory (Walling and Harley, 2004). A. Neuronal Development NE plays a role in modulating the development of neurons in various brain areas. For example, in rat visual cortex NE deafferentation leads to various changes, including the impairment of stimulus selectivity and reduction of the signal-to-noise ratio. In contrast, ACh deafferentation leads only to a transient disturbance of ocular dominance (Siciliano et al., 1999). In embryonic rat brain, local delivery of NE in the neocortex regulates the survival and proliferation of cells in the germinal epithelium. Lesions that interfere with this process lead to apoptosis (Pabbathi et al., 1997; Popovik and Haynes, 2000). NE exerts these effects by activating Ars, thus raising cAMP levels. This promotes the expression of c-fos and induction of the enzyme ornithine decarboxylase that is obligatory for neuronal development. Interference with this system at critical stages of development leads to lasting changes in neuronal differentiation and responsiveness (Wagner et al., 1995). Happe et al. (2004) review the complex role of -2 receptors in brain development. B. Stress Stress of various kinds is associated with activation of the NE system. For example, stress (foot shock and restraint) increases NE, DA, and 5-HT metabolism in the cortex, HC, and brainstem; this effect is not mediated by NO (Dunn, 1998). However, Ishizuka et al. (2000) found that 20 minutes of intermittent foot shock in rats led to increased levels of NE in the HT PVN and medial PFC. This was associated with increased levels of NO in the PFC but not in the PVN. Stress induces increases in e.c. NE and DA in rat medial PFC in complex ways that are

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differentially affected by the tranquilizer diazepam (Finlay et al., 1995). Kawahara et al. (2000) confirm that handling stress promotes NE release in the medial PFC in rats. They addressed the question of which afferents to the LC mediate this response but were successful only in eliminating glu, GABA, and ACh systems. Stress also raises NE levels in rat HC (Petty et al., 1993). In contrast, Sajdyk et al. (1997) found that only the full fear-potentiated startle stress test, and not foot shocks or acoustic startle, led to increased NE and DA levels in the dorsomedial hypothalamus. Another study showed that stress by itself (manual restraint) led to increased NE release in rat PVN that rapidly attenuated when the stress was repeated every 120 minutes. But manual restraint plus tail pinch led to a greater degree of NE release that did not attenuate (Shibaski et al., 1995). In contrast, Nakane et al. (1994) reported that immobilization stress by itself activated NE release in rat PFC that did not attenuate for the period of the stress nor for 20 minutes afterwards. Fulford and Marsden (1997) reared rats in social isolation for 6 to 8 weeks. Microdialysis measurement showed that this procedure induced enhancement of presynaptic 2 ARs in the dorsal HC that resulted in decreased functional responsiveness of HC NE nerve terminals. Both single and repeated immobilization stress induced long-lasting (24 hours) increases in mRNA levels in the A5 NE cell group (Micutkova et al., 2001). Shaker stress induces raised NE levels in the HT in rats. This effect is attenuated by NO donors (Hashiguchi et al., 1997). NE also modulates the synthesis of GABA in central limbic stress circuits. The antidepressant reboxetine is a selective NE reuptake blocker and inhibits glutamic acid decarboxylase, the key enzyme in GABA synthesis, in various limbic brain areas (Herman et al., 2003). Fear-related stimuli activate oxytocin neurons in the HT and facilitate oxytocin release in the pituitary. This effect is mediated by the NE system. An unconditioned fear stimulus induces the expression of Fos protein in magnocellular oxytocin neurons in the SON and PVN. The same stimulus induces the expression of Fos protein in both A1 and A2 NE neurons and in C1 and C2 adrenergic neurons. Conditioned-fear stimuli induce Fos expression preferentially in A2/C2 neurons (Zhu and Onaka, 2002). Acute psychological stress increases the number of blood natural killer cells associated curiously with a significantly reduced production of ROS (Atanackovic et al., 2002). C. Behavior The NE system plays a prominent role in brain mechanism subserving alertness and attention. Coull et al. (1999) carried out PET studies in normal humans and showed that NE mediates functional integration between various


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attentional systems in the brain. In particular, it potentiates connectivity between the LC ! parietal cortex and between parietal cortex ! thalamus and frontal cortex. In rat experiments, Dalley et al. (2001) reported that the NE system is specifically involved in detecting shifts in the predictive relationship between instrumental action and reinforcement. Thus, NE can be said to be involved in the detection of novel task contingencies. In contrast, ACh pays a crucial role in the detection of visual targets. Attentional level is modulated positively by the amount of NE released, whereas attentional quality depends on the interaction of the release of glu and the release of NE (Kodama et al., 2002). Stimulation of presynaptic 2 ARs in the medial PON effects an arousal response, whereas stimulation of postsynaptic 1 ARs in this nucleus induces sleep (Ramesh and Kumar, 1998). Mallick and Joseph (1998) present evidence to suggest that the interaction and integration between the NE and ACh inputs to the medial POA modulate sleeping/waking. During slow-wave sleep LC, neurons in the rat decrease their activity and become virtually quiescent during REM sleep. This is due to an inhibitory GABAergic tone (Gervasoni et al., 1998). Nitz and Siegel (1997) confirm in the cat that inhibitory GABAergic tone is responsible for the cessation of discharge of LC neurons during REM sleep. There are two types of cholinergic neuron in the LTDN that play key roles in arousal. Type 1-S neurons exhibit a slow tonic discharge during both waking and REM sleep. Type PGO-on neurons produce single spikes during waking and burst discharges and PGO waves during REM sleep (Koyama and Sakai, 2000). NE and E (as well as glu and histamine) induce tonic discharges in both types of neuron. Carbachol inhibited single spikes in both types but had no effect on PGO-on burst discharges. DA had no effect on either type of neuron (Koyama and Sakai, 2000). The transition from sleep to waking is associated with abolition of spindle waves in thalamocortical neurons and in the reticular nucleus of the thalamus. Local applications of NE and 5-HT abolish spindle waves by enhancing the hyperpolarization-activated cation current Ih in thalamocortical neurons (Lee and McCormick, 1996). Hajnal et al. (1997) injected NE into the PVN and monitored by microdialysis e.c. levels of NE and ACh in the nucleus accumbens. NE injections effectively induced eating behavior in satiated rats. In a separate test, when the same animals were tested in the absence of food, NE injections at the same dosage caused a significant rise in e.c. DA and a decrease in e.c. ACh levels. In contrast, when food was available and ingested, ACh levels increased instead of decreasing. The authors suggest that these results indicate that functional links exist between the PVN and NAc in which DA helps to initiate, and ACh helps stop, appetitive behavior involved in the reinforcement of eating (Hajnal et al., 1997).

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The NE system also plays a role in various types of emotional behavior such as irritability and hostility. A genetic study of two sets of human subjects showed that variations in the 2A AR gene were significantly correlated with increased hostility, irritability, negativity, and verbal aggression (Comings et al., 2000). In rat brain slices, NE application to neurons in the periaqueductal gray (PAG) depolarized most neurons in the ventrolateral portion but had a mixed response (equal proportions of depolarization and hyperpolarization) in the lateral portion. Thus, in the PAG, NE biases activity in favor of a ventrolateral-mediated response pattern that includes quiescence, hyporeactivity, hypotension, and bradycardia (Vaughan et al., 1996).

VIII. Miscellaneous Items

A. Alzheimer’s Disease NE mechanisms are also involved in the complex series of biochemical changes associated with AD. NE, in general, inhibits inflammatory gene expression. Degeneration of the NE neurons in the LC is a well-established neuropathological feature of AD. In experimental animals, NE depletion potentiates the inflammatory response to A with increases in iNOS, IL-1, and IL-6 expression (Heneka et al., 2002). -amyloid (1–40) (A) also inhibited by 50% the electrically stimulated release of NE in rat cortex slices and increased i.c. Ca2þ levels by 78%. Both these effects were mediated by ROS, being prevented by ROS scavengers (Li and Smith, 1996). Fu et al (1998) found that catecholamines (but not 5-HT nor ACh) potentiated A toxicity to cultured HC neurons by inducing oxidative stress. Catecholamines and A synergistically caused a decrease in mitochondrial transmembrane potential and function. A also induced a slow increase in i.c. Ca2þ levels, which was much exacerbated by catecholamines. Levels of membrane lipid peroxidation and cytoplasmic and mitochondrial ROS were all increased by A. These indices of oxidative stress were also exacerbated by catecholamines (Fu et al., 1998). B. Anticholinesterase Neurotoxicity The main toxic effects of anticholinesterase nerve gases such as sarin and soman are exerted by excess cholinergic activity. However, these agents also affect the NE system. Soman inhibits NMDA-stimulated NE release by interfering with some of the regulatory sites on the NMASR/ion channel complex. It also interferes with voltage-dependent Naþ channel function, but not with


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VOCCs (Tang and Cassel, 1998). In rats, soman induces a complex series of events over time. As early as 3 minutes after the onset of soman-induced seizures, cortical levels of ACh rose. Then after 5 minutes, NE levels fell, and DA levels rose. By 20 minutes, levels of aspartate and glutamate fell, and levels of GABA rose (Shih and McDonough, 1997). C. Gap Junctions NE also modulates the activity at gap junctions between astrocytes and neurons in rat striatal cultures. NE stimulates ,1 ARs leading to a depression of the permeability of the gap junction. This effect is mainly mediated by the 1 AR ! PLC ! IP ! PKC cascade and partly by the AR ! cAMP cascade (Giaume et al., 1991). D. Oxytocin (OT)/Vasopressin (VP) In rat PVN, magnocellular neurons receive excitatory inputs from local intranuclear glu neurons. The latter are activated by NE by means of 1 ARs (Daftary et al., 1998). In mouse HT, NE activates VP and OT expression in both the PVN and the SON, whereas 5-HT activates VP and OT expression in the PVA, but only OT expression in the SON (Vacher et al., 2002). In the other direction, VP potentiates the NE-induced stimulation of cAMP formation in both neurons and glia in the HC (Brinton et al., 2000). E. Opioids Opioid peptides (including -endorphin) inhibit Kþ-stimulated NE release in the caudal dorsomedial medulla; Carr and Gregg, 1995). Presynaptic -opioid receptors inhibit Kþ-evoked NE release in rat HC (Matsumoto et al., 1994). F. Transporters The activity of NE and DA in synapses in the brain is largely terminated by MAO or by rapid uptake into presynaptic terminals by high-affinity Naþ/Cl–– dependent transporters. COMT is not normally significant for the inactivation of catecholamines but becomes so under circumstances when catecholamine synthesis is increased (Huotari et al., 2002). The NE-dependent transporter is also subject to endocytosis and rapid recycling. Second messengers in this system

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include protein kinases and protein phosphatases, in particular PKC, as well as AA, ROS, and NO (Zahniser and Doolen, 2001). In PC-12 cells Ca2þdependent enhancement of NE uptake by the NE transporter is mediated by activation of calmodulin-dependent protein kinases. This probably involves stimulation of recycling of the NE transporter to the plasma membrane and/or direct phosphorylation of the transporter molecule (Uchida et al., 1998). All NE neurons in the LC and the adjacent lateral tegmentum express mRNA for the NE transporter. It is interesting that brainstem E neurons do not contain mRNA for the NE transporter. This may indicate that E uses its own transporter or that E may act as an endocrine regulator that does not require a rapid reuptake mechanism (Lorang et al., 1994).

G. Histamine In mouse and guinea-pig cortex, activation of presynaptic histamine H3R1 receptors inhibits NE release, whereas activation of nonpresynaptic H2Rs augments NE release (Timm et al., 1998).

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Venkatesan, C., Song, X.-Z., Go, C.-G., Kurose, H., and Aoki, C. (1996). Cellular and subcellular distribution of 2A-adrenergic receptors in the visual cortex of neonatal and adult rats. J. Comp. Neurol. 365, 79–95. Wagner, E. J., Ronnekleiv, O. K., and Kelly, M. J. (2001). The noradrenergic inhibition of an apamin-sensitive, small-conductance Ca2þ-activated Kþ channel in hypothalamic gammaaminobutyric acid neurons: Pharmacology, estrogen sensitivity, and relevance to the control of the reproductive axis. J. Pharmacol. Exp. Ther. 299, 21–30. Walaas, S. I. (1999). Regulation of calcium-dependent [3H]noradrenaline release from rat cerebrocortical synaptosomes by protein kinase C and modulation of the actin cytoskeleton. Neurochem. Int. 34, 221–233. Walling, S. G., and Harley, C. W. (2004). Locus ceruleus activation initiates delayed synaptic potentiation of perforant path input to the dentate gyrus in awake rats: A novel betaadrenergic- and protein synthesis-dependent mammalian plasticity mechanism. J. Neurosci. 24, 598–604. Wang, S. J., Cheng, L. L., and Gean, P. W. (1999). Cross-modulation of synaptic plasticity by betaadrenergic and 5-HT1A receptors in the rat basolateral amygdala. J. Neurosci. 19, 570–577. Wang, Y. F., Shibuya, I., Kabashima, N., Setiadji, V. S., Isse, T., Ueta, Y., and Yamashita, H. (1998). Inhibition of spontaneous inhibitory postsynaptic currents (IPSC) by noradrenaline in rat supraoptic neurons through presynaptic alpha2-adrenoceptors. Brain Res. 807, 61–69. Warren, R. A., and Dykes, R. W. (1996). Transient and long-lasting effects of iontophoretically administered norepinephrine on somatosensory cortical neurons in halothane-anesthetized cats. Can. J. Physiol. Pharmacol. 74, 38–57. Williams, J. A., and Reiner, P. B. (1993). Noradrenaline hyperpolarizes identified rat mesopontine cholinergic neurons in vitro. J. Neurosci. 13, 3878–3883. Xin, W., Shen, X. M., Li, H., and Dryhurst, G. (2000). Oxidative metabolites of 5-Scysteinylnorepinephrine are irreversible inhibitors of mitochondrial complex I and the alphaketoglutarate dehydrogenase and pyruvate dehydrogenase complexes: Possible implications for neurodegenerative brain disorders. Chem. Res. Toxicol. 13, 749–760. Yamada, H., Hayashi, M., Uehara, S., Kinoshita, M., Muroyama, A., Watanabe, M., Takei, K., and Moriyama, Y. (2002). Norepinephrine triggers Ca2þ-dependent exocytosis of 5-hydroxytryptamine from rat pinealocytes in culture. J. Neurochem. 81, 533–540. Yamada, H., Yamamoto, A., Yodozawa, S., Kozaki, S., Takahashi, M., Morita, M., Michibata, H., Furuichi, T., Mikoshiba, K., and Moriyama, Y. (1996). Microvesicle-mediated exocytosis of glutamate is a novel paracrine-like chemical transduction mechanism and inhibits melatonin secretion in rat pinealocytes. J. Pineal Res. 21, 175–191. Yamashita, H., Sato, N., Kizaki, T., Oh-ishi, S., Segawa, M., Saitoh, D., Ohira, Y., and Ohno, H. (1995). Norepinephrine stimulates the expression of fibroblast growth factor 2 in rat brown adipocyte primary culture. Cell Growth Differ. 6, 1457–1462. Yang, B., Wang, Y., and Cynader, M. S. (1996). Synergistic interactions between noradrenaline and glutamate in cytosolic calcium influx in cultured visual cortical neurons. Brain Res. 721, 181–190. Yang, H. W., Lin, Y. W., Yen, C. D., and Min, M. Y. (2002). Change in bi-directional plasticity at CA1 synapses in hippocampal slices taken from 6-hydroxydopamine-treated rats: The role of endogenous norepinephrine. Eur. J. Neurosci. 16, 1117–1128. Yokotani, K., Wang, M., Murakami, Y., Okada, S., and Hirata, M. (2000). Brain phospholipase A(2)arachidonic acid cascade is involved in the activation of central sympatho-adrenomedullary outflow in rats. Eur. J. Pharmacol. 398, 341–347. Yuan, P. X., Huang, L. D., Jiang, Y. M., Gutkind, J. S., Manji, H. K., and Chen, G. (2001). The mood stabilizer valproic acid activates mitogen-activated protein kinases and promotes neurite growth. J. Biol. Chem. 276, 31674–31683.


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Yunker, W. K., Lee, E. K., Wong, A. O., and Chang, J. P. (2000). Norepinephrine regulation of growth hormone release from goldfish pituitary cells. II. Intracellular sites of action. J. Neuroendocrinol. 12, 323–333. Zahniser, N. R., and Doolen, S. (2001). Chronic and acute regulation of Naþ/Cl- -dependent neurotransmitter transporters: Drugs, substrates, presynaptic receptors, and signaling systems. Pharmacol. Ther. 92, 21–55. Zhang, C., Guo, Y. Q., Qiao, J. T., and Dafny, N. (1998). Locus coeruleus modulates thalamic nociceptive responses via adrenoceptors. Brain Res. 784, 116–122. Zhu, L., and Onaka, T. (2002). Involvement of medullary A2 noradrenergic neurons in the activation of oxytocin neurons after conditioned fear stimuli. J. Neurosci. 16, 2186–2198.

Further Reading

Castro-Alamancos, M. A. (2002). Properties of primary sensory (lemniscal) synapses in the ventrobasal thalamus and the relay of high-frequency sensory inputs. J. Neurophysiol. 87, 946–953. Ferraro, L., Tanganelli, S., Calo, G., and Antonelli, T. Noradrenergic modulation of gammaaminobutyric acid outflow from the human cerebral cortex. Brain Res. 629, 103–108. Kalsner, S., and Abdali, S. A. (2002). Rate-independent inhibition of 5-HT release by 5-HT in the somatodendritic regions of raphe neurons. Brain Res. Bull. 58, 289–294. Matsumoto, M., Yoshioka, M., Togashi, H., Tochihara, M., Ikeda, T., and Saito, H. (1995). Modulation of norepinephrine release by serotonergic receptors in the rat hippocampus as measured by in vivo microdialysis. J. Pharmacol. Exp. Ther. 272, 1044–1051. Pisani, A., Bonsi, P., Centonze, D., Martorana, A., Fusco, F., Sancesario, G., De Persis, C., Bernardi, G., and Calabresi, P. (2003). Activation of beta1-adrenoceptors excites striatal cholinergic interneurons through a cAMP-dependent, protein kinase-independent pathway. J. Neurosci. 23, 5272–5282. Sershen, H., Balla, A., Lajtha, A., and Vizi, E. S. (1997). Characterization of nicotinic receptors involved in the release of noradrenaline from the hippocampus. Neuroscience 77, 121–130. Mooney, E. E., and Tavassoli, F. A. (1999). Papillary transitional cell carcinoma of the breast: A report of five cases with distinction from eccrine acrospiroma. Mod. Pathol. 12, 287–294. Troadec, J. D., Marien, M., Darios, F., Hartmann, A., Ruberg, M., Colpaert, F., and Michel, P. P. (2001). Noradrenaline provides long-term protection to dopaminergic neurons by reducing oxidative stress. J. Neurochem. 79, 200–210. Ivon Ku¨ gelgen, I., and Starke, K. (1995). Kainate receptors are involved in the glutamate-induced indirect, purinergic inhibition of [3H]-noradrenaline release in rabbit brain cortex. Naunyn Schmiedebergs Arch. Pharmacol. 353, 46–54.

SECTION IV THE ADRENALINE SYSTEM

The adrenergic system in the brain has attracted much less attention than its fellow catecholamines dopamine and norepinephrine. The adrenaline-containing neurons in the brain are restricted to three small groups in the medulla labeled C1, C2, and C3. Ruggiero et al. (1985) describe the detailed anatomy in the rat. Some of these neurons project downward to the spinal cord, where they modulate sympathetic neurons. Others project rostrally, where they innervate a number of key structures. The C1 group project strongly to the SN (Nagatsu et al., 1998: mouse); the hypothalamus (Verberne et al., 1999: rat); the locus coeruleus (inhibitory) (Aston-Jones et al., 1992: rat; Pieribone and Aston-Jones 1991: rat; Pieribone et al., 1988: rat); the central nucleus of the amygdala (many of these neurons also contain neuropeptide Y) (Roder and Ciriello, 1993: rat; Zardetto-Smith and Gray, 1995: rat); the raphe nucleus (Luppi et al., 1989: cat); the paraventricular and supraoptic nuclei of the hypothalamus (Cunningham et al., 1990: rat); the central tegmentum (Haselton and Guyenet, 1989: rat); the periaqueductal gray (Herbert and Saper, 1992: rat); and sparsely to the REM center in the dorsolateral mesopontine tegmentum (Semba, 1993: rat). Adrenergic aVerents from the medullary nuclei modulate activity in the midline paraventricular nucleus of the thalamus by two disynaptic pathways. The first (lateral) runs through the central nucleus of the amygdala, sublenticular substantia innominata, and bed nucleus of the stria terminalis. The second (medial) includes the lateral hypothalamic area, the perifornical nucleus, and dorsomedial and periventricular hypothalamic nuclei (Otake et al., 1995: rat). The C1 and C3 groups project to the dorsal raphe (Peyron et al., 1996: rat). The C1, C2, and C3 groups all project strongly to the histaminic neurons of the tuberomammillary nucleus of the posterior hypothalamus (Ericson et al., 1989: rat) and to the paraventricular nucleus of the hypothalamus (Phillipson and Bohn, 1994: rat; Sawchenko and Bohn, 1989: rat). Rostral input to these nuclei includes axons from the nucleus of the tractus solitarius that mediates sympathetic stimulation (Aicher et al., 1996 and from the central nucleus of the amygdala (Cassell and Gray, 1989: rat). This system is certainly concerned with the control of various lowlevel autonomic functions such as blood pressure, heart rate, and respiration (Ceccatelli et al., 1989; Reis et al., 1989). However, the wide nature and extent of its rostral projections to key limbic structures suggests that it may play a yet undiscovered role in higher mental functions. Further research in this area is urgently needed.


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References

Aicher, S. A., Saravay, R. H., Cravo, S., Jeske, I., Morrison, S. F., Reis, D. J., and Milner, T. A. (1996). Monosynaptic projections from the nucleus tractus solitarii to C1 adrenergic neurons in the rostral ventrolateral medulla: Comparison with input from the caudal ventrolateral medulla. J. Comp. Neurol. 373, 62–75. Aston-Jones, G., Astier, B., and Ennis, M. (1992). Inhibition of noradrenergic locus coeruleus neurons by C1 adrenergic cells in the rostral ventral medulla. Neuroscience 48, 371–381. Cassell, M. D., and Gray, T. S. (1989). The amygdala directly innervates adrenergic (C1) neurons in the ventrolateral medulla in the rat. Neurosci. Lett. 97, 163–168. Ceccatelli, S., Millhorn, D. E., Hokfelt, T., and Goldstein, M. (1989). Evidence for the occurrence of an enkephalin-like peptide in adrenaline and noradrenaline neurons of the rat medulla oblongata. Exp. Brain Res. 74, 631–640. Cunningham, E. T., Jr., Bohn, M. C., and Sawchenko, P. E. (1990). Organization of adrenergic inputs to the paraventricular and supraoptic nuclei of the hypothalamus in the rat. J. Comp. Neurol. 292, 651–667. Ericson, H., Blomqvist, A., and Kohler, C. (1989). Brainstem aVerents to the tuberomammillary nucleus in the rat brain with special reference to monoaminergic innervation. J. Comp. Neurol. 281, 169–192. Haselton, J. R., and Guyenet, P. G. (1989). Electrophysiological characterization of putative C1 adrenergic neurons in the rat. Neuroscience 30, 199–214. Herbert, H., and Saper, C. B. (1992). Organization of medullary adrenergic and noradrenergic projections to the periaqueductal gray matter in the rat. J. Comp. Neurol. 315, 34–52. Luppi, P. H., Fort, P., Kitahama, K., Denoroy, L., and Jouvet, M. (1989). Adrenergic input from medullary ventrolateral C1 cells to the nucleus raphe pallidus of the cat, as demonstrated by a double immunostaining technique. Neurosci. Lett. 106, 29–35. Nagatsu, I., Ikemoto, K., Takeuchi, T., Arai, R., Karasawa, N., Fujii, T., and Nagatsu, T. (1998). Phenylethanolamine-N-methyltransferase—immunoreactive nerve terminals aVerent to the mouse substantia nigra. Neurosci. Lett. 245, 41–44. Otake, K., Ruggiero, D. A., and Nakamura, Y. (1995). Adrenergic innervation of forebrain neurons that project to the paraventricular thalamic nucleus in the rat. Brain Res. 697, 17–26. Peyron, C., Luppi, P. H., Fort, P., Rampon, C., and Jouvet, M. (1996). Lower brainstem catecholamine aVerents to the rat dorsal raphe nucleus. J. Comp. Neurol. 364, 402–413. Phillipson, O. T., and Bohn, M. C. (1994). C1–3 adrenergic medullary neurones project to the paraventricular thalamic nucleus in the rat. Neurosci. Lett. 176, 67–70. Pieribone, V. A., and Aston-Jones, G. (1991). Adrenergic innervation of the rat nucleus locus coeruleus arises predominantly from the C1 adrenergic cell group in the rostral medulla. Neuroscience 41, 525–542. Pieribone, V. A., Aston-Jones, G., and Bohn, M. C. (1988). Adrenergic and non-adrenergic neurons in the C1 and C3 areas project to locus coeruleus: A fluorescent double labeling study. Neurosci. Lett. 85, 297–303. Reis, D. J., Ruggiero, D. A., and Morrison, S. F. (1989). The C1 area of the rostral ventrolateral medulla oblongata. A critical brainstem region for control of resting and reflex integration of arterial pressure. Am. J. Hypertens. 2, 363S–374S. Roder, S., and Ciriello, J. (1993). Innervation of the amygdaloid complex by catecholaminergic cell groups of the ventrolateral medulla. J. Comp. Neurol. 332, 105–122. Ruggiero, D. A., Ross, C. A., Anwar, M., Park, D. H., Joh, T. H., and Reis, D. J. (1985). Distribution of neurons containing phenylethanolamine N-methyltransferase in medulla and hypothalamus of rat. J. Comp. Neurol. 239, 127–154.

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Sawchenko, P. E., and Bohn, M. C. (1989). Glucocorticoid receptor-immunoreactivity in C1, C2, and C3 adrenergic neurons that project to the hypothalamus or to the spinal cord in the rat. J. Comp. Neurol. 285, 107–116. Semba, K. (1993). Aminergic and cholinergic aVerents to REM sleep induction regions of the pontine reticular formation in the rat. J. Comp. Neurol. 330, 543–556. Verberne, A. J., Stornetta, R. L., and Guyenet, P. G. (1999). Properties of C1 and other ventrolateral medullary neurones with hypothalamic projections in the rat. J. Physiol. 517, 477–494. Zardetto-Smith, A. M., and Gray, T. S. (1995). Catecholamine and NPY eVerents from the ventrolateral medulla to the amygdala in the rat. Brain Res. Bull. 38, 253–260.

SECTION V SEROTONIN SYSTEM

I. Anatomy A. 5-HT1 Receptors B. 5-HT2 Receptors C. Other or Unspecified 5-HT Receptors D. Synaptic Versus Volume Transmission II. Electrical Actions A. Potassium Related B. Role of Calcium Channels III. Interactions Between Serotonin and Other Neurotransmitter Systems A. Actions of Serotonin on Glutamate Synaptic Transmission B. Actions of Glutamate on Serotonin Systems C. Actions of Serotonin on GABA Systems D. Actions of GABA on Serotonin Systems E. Actions of Serotonin on the Cholinergic System F. Actions of Serotonin on the Dopamine System G. Actions of Serotonin on the Norepinephrine System H. EVects of Serotonin on the Nitric Oxide System I. EVects of Nitric Oxide on the Serotonin System IV. Postsynaptic Cascades A. 5-HT(1A)R B. 5-HT(1B)R C. 5-HT(1D)R D. 5-HT(1E)R E. 5-HT(1F)R F. 5-HT(2A)R G. 5-HT(2B)R H. 5-HT(2C)R (Confined to Nervous Tissue) I. 5-HT(4)R (Mostly in Heart and Gut) J. 5-HT(5)R (Confined to Nervous Tissue) K. 5-HT(6)R (Predominantly in Brain) L. 5-HT(7)R (Plentiful in Brain) V. Serotonin and Synaptic Plasticity A. Serotonin and ScaVolding Proteins B. Serotonin and Learning C. Stress D. Anxiety VI. Serotonin and the Development of Neural Systems VII. Sleep VIII. Miscellaneous A. Interleukins B. Estrogens C. Opioids D. Melatonin E. Cannabinoids


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F. Neurokinin G. Lysergic Acid Diethylamide (LSD) H. Reactive Oxygen Species (ROS) I. Apoptosis J. ERK K. Hormones L. Peptides M. Endocytosis N. Aging O. Dementia P. Social Dominance Behavior Q. Prepulse Inhibition R. Beauty and Spirituality References

I. Anatomy

Serotonin and serotonin-like chemicals appeared in the evolution of biological systems very early on and played an important role in light-processing biochemistry in plants, where they direct growth toward the source of light. In animals, serotonin modulates the cytoskeleton to influence the formation cellular contacts and aVects the proliferation, migration, and maturation of many cell types (Azmitia, 2001). The brain serotonin (5-HT) system influences a wide range of functions, including autonomic, circadian rhythms, appetite, aggression, sensorimotor activity, sexual behavior, mood, cognition, perception, learning, memory, and personality. As Daw et al. (2002) say, ‘‘Serotonin plays an extremely complicated set of roles in the brain, roles that it is impossible at present to encompass within a single theory.’’ So a piecemeal approach is the best we can do. The principal raphe nuclei in the brainstem are the dorsal nucleus, which projects mainly by thin fibers to the cerebral cortex, the median nucleus, which projects by thick fibers with large varicosities to the hippocampus, and the caudal nuclei, which project to the spinal cord (Kosofsky and Molliver, 1987). The raphe 5-HT system innervates practically all the brain and is the first neurotransmitter system to develop in the embryo. Serotonin is an important cellular chemical, even very low down in the phylogenetic chain. In plants, its light-adsorbing properties play an important role in photosynthesis. Serotonin modulates, in particular, neurogenesis, apoptosis, dendritic and spine refinement, cell migration, and synaptic plasticity (Sodhi and Bush, 2004). Growth factors for 5-HT neurons include S100, BDNF, Petl transforming-growth factor bets, bone morphogenetic factor, and ciliary neurotrophic factor (Sodhi and SandersBush, 2004).

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Leger et al. (2001) provide a detailed account of the distribution of 5-HT cell bodies and axons in the cat brain. The cell bodies are located mainly in the raphe nuclei, but some are also found in the ventrolateral and dorsal reticular formation. Axons are seen in every region of the gray matter, except the superior olivary complex. They are particularly common in the olfactory tubercle, nucleus accumbens, ventral mesencephalon, periventricular gray, and inferior olive. In the neocortex 5-HT axons form tight varicose pericellular arrays. Further details of the serotoninergic projection from the raphe nuclei to the anterior thalamic nuclei are given by Gonzalo-Ruiz et al. (1995). Vertes et al. (1999) present a detailed account of the projections of the median nucleus in the rat. The dorsal and median nuclei project to nonoverlapping areas of forebrain. The latter projects mainly to midline structures, with a few projections to the cortex (perirhinal, entorhinal, and frontal); the latter projects widely to the cortex. Smith (1999) states that the thalamic midline and dorsal nuclei contain high levels of serotonin receptors in several species. The raphe nuclei are chemically highly heterogenous. 5-HT neurons throughout show distinct morphologies and densities in each subdivision of the raphe (median, lateral, dorsal, ventrolateral, ventral, and caudal). There are also many SP-, GABA-, and PV-neurons in each subdivision plus some scattered THþ, calbindinþ, and calretininþ cells (Charara and Parent, 1998). In human postmortem studies, Oke et al. (1997) found 5-HT to be present in medial and intralaminar thalamic nuclei and in the pulvinar. Li et al. (2001) report that 5-HT neurons in the dorsal raphe have dendritic spines, whereas non-5-HT neurons in this location do not. In the rat septum, 5-HT terminals form two groups (1) symmetrical synapses on somata and proximal dendrites and (2) asymmetrical synapses on small distal dendrites of unidentified neurons (Milner and Veznedaroglu, 1993). In rat SNpc, 5-HT terminals are equally distributed between synaptic contacts on dendrites and nonsynaptic (volume) locations. In contrast, in the SNpr, most 5-HT terminals are synaptic on DA and GABAergic neurons (Moukhles et al., 1997). In rat bed nucleus of the stria terminalis, 5-HT axons form synapses only in the ventrolateral region. In contrast, DA terminals synapse on dendrites, spines, somata, and axons mainly in the lateral region, and NE terminals synapse mainly on dendrites and spines in the medial region. There are also dense networks of NE terminals in the intermediate and some in the dorsolateral region (Phelix et al., 1992). Gao and Mason (1997) give a detailed account of the microanatomy of the 5-HT neurons in the rat medulla. Gonzales-Ruiz et al. (1995) do the same for the projections from the raphe nuclei to the anterior thalamic nuclei in the rat. Losier and Semba (1993) found that some 4.7% of 5-HT neurons in the dorsal, median, and caudal linear raphe nuclei have dual projections both to the

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thalamus and to the basal forebrain. The latter represents the major cholinergic input to the cerebral cortex. Likewise, 8% of cholinergic neurons in the PPN and LDTN and 1.5 % of NE in the locus ceruleus also exhibit this type of dual projection. Thus, this system allows conjoint modulation of the activity of neurons in the thalamus and cortex during cortical arousal (Losier and Semba, 1993). A. 5-HT1 Receptors Levels of the 5-HT(1A)R in living human brain are high in the cerebral cortex, hippocampus, and raphe and are low in the thalamus and basal ganglia (Ito et al., 1999). Sanabrier-Bohorquez et al. (2002) reported from human PET studies that the number of 5-HT(1A)Rs is ranked HC > neocortex > basal ganglia and cerebellum. In rat cerebral cortex, HC, hypothalamus, and amygdala 5-HT(1A)Rs are located on principal cells, calbindinþ and parvalbuminþ interneurons (INS); in the lateral septum and thalamus, they occur on INS (both types) only; and in the medial septum and reticular nucleus of the thalamus on parvalbuminþ INs only (Aznar et al., 2003). In the medial PFC, pyramidal neurons colocalize 5-HT(1A) and 5-HT(2A) Rs (Martin-Ruiz et al., 2001). In rat raphe, 5-HT(1A)Rs are located only postsynaptically on somata and on dendrites mostly extrasynaptically. In the HC, these receptors are located on spines, dendrites, and somata of pyramidal and granule cells. In both SN and globus pallidus, 5-HT(1B)Rs are located mainly on fine preterminal axons but not on the actual terminals (Riad et al., 2000). In addition, throughout the brain, many 5-HT(1A)Rs are inhibitory presynaptic autoreceptors exerting negative feedback on serotoninergic somata and dendrites (Blier et al., 1998). In the parabrachial and paranigral subdivisions of the VTA in rat brain, 5-HT(1A)Rs are mostly postsynaptic on the soma and dendrites of DA cells, with a few presynaptic and a few on glia. Many are located intracellularly in the smooth endoplasmic reticulum. The authors suggest that the latter represent a reserve of receptors susceptible to mobilization to functional postsynaptic sites on the plasma membrane by aVerent input to distal dendrites in the VTA (Doherty and Pickel, 2001). The distribution of the mRNAs for the 5-HT(1B)Rs and the 5-HT(ID)R in guinea-pig brain is detailed by Bonaventure et al. (1998). The former had a wide distribution throughout the brain, but the latter were restricted mainly to the olfactory tubercle, entorhinal cortex, dorsal raphe, cerebellum, mesencephalic trigeminal nucleus, and the trigeminal ganglion. The authors suggest that their finding that the 5-HT1B receptor exerts an important part of its influence in the striatum at the presynaptic level (e.g., on cortical axon endings) would imply that,

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within the basal ganglia, the receptor is involved in presynaptic regulation of both excitatory (glutamate) and inhibitory (GABA) neurotransmitters. The 5-HT (1D)R was particularly plentiful in the nucleus of the trigeminal nerve. This suggested that it might play a role in the release of inflammatory polypeptides such as substance P. In the mouse, 5-HT(1B)Rs are located mainly in the basal ganglia and are mostly presynaptic on axon terminals (Boschert et al., 1994). The presynaptic 5-HT autoreceptors that modulate 5-HT release in rat ventral LGN are of the 1B and 1D subtypes (Davidson and Stamford, 1996). Inhibitory 5-HT(1D) autoreceptors are presynaptic on terminals in the guinea-pig cerebral cortex (Buhlen et al., 1996; Roberts et al., 1996). Sixty percent of the neurons in rat and mouse prefrontal cortex express 5-HT1A and/or 5-HT2A receptor mRNAs, which are highly colocalized (80%) (Amargos-Bosch et al., 2004). Electrical stimulation of the dorsal and median raphe nuclei elicits 5-HT1A– mediated inhibitions and 5-HT2A–mediated excitations in identified pyramidal neurons recorded extracellularly in rat medial prefrontal cortex (mPFC). These authors found that opposite responses in the same pyramidal neuron could be evoked by stimulating the raphe nuclei at diVerent coordinates. This finding suggested a precise connectivity between 5-HT neuronal subgroups and 5-HT1A and 5-HT2A receptors in pyramidal neurons. B. 5-HT2 Receptors In monkey cortex, 5-HT(2A)Rs are located both presynaptically and postsynaptically in all cortical layers, especially in layers II, III, V, and VI, with only a few in layer IV. These are located mainly on pyramidal cells, particularly on their apical dendrites, on INS, and on axon terminals (Jakab and Goldman-Rakic, 1998). In human brain, these receptors are located quantitatively in the cerebral cortex > HC, hypothalamus, nucleus accumbens, amygdala >> thalamus, caudate, putamen, and cerebellum (Dwivedi and Pandey, 1998). In postmortem human brain, Hall et al. (2000) reported many of these receptors in the cortex; few in the HC, basal ganglia, and thalamus; and none in the cerebellum and brainstem. In rat frontal cortex (layer V), 5-HT(2A)Rs are found on the somata and dendrites of pyramidal cells, as well as in the HC, amygdala, striatum, olfactory structures, and on astrocytes (Xu and Pandey, 2000). In rat motor striatum (caudate/putamen) and limbic striatum (N.Ac.), 5-HT (2A)Rs are located mainly on dendrites (but not spines), with some on somata and axons. Most of these are located intracellularly in association with cytoplasmic organelles. There is also some colocalization of 5-HT(2A)Rs with NMDAR1s (Rodriguez et al., 1999). Fay and Kubin (2000) give a very detailed account of the distribution of 5-HT(2A)Rs in rat pons and medulla. In the rat PPN/LDTN,

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these receptors are located on INS but not on cholinergic main neurons (Fay and Kubin, 2001). In rat prelimbic PFC of the 5-HT(2A)R population, 73% are postsynaptic on dendrites and spines of pyramidal cells and dendrites only of INs, whereas 24% are presynaptic located mainly on aminergic axons (Miner et al., 2003). In the striatum, 5-HT(2A)Rs are heteroreceptors on fibers coming from the cortex and pallidum to synapse on parvalbuminþ INS (Bubser et al., 2001). 5-HT(2C)Rs are mainly postsynaptic in various brain areas in the rat, especially in the midbrain and brainstem (Sharma et al., 1997). Clemett et al. (2000) found that the areas containing most 5-HT(3)Rs in rat brain were anterior olfactory nucleus, medial and intercalated amygdala nuclei, hippocampal Ca1 and CA3, raphe, laterodorsal and lateral geniculate nuclei, caudate/putamen, and several cortical areas (including piriform and frontal). C. Other or Unspecified 5-HT Receptors In the rat, mRNA for the 5-HT(7)R is located particularly densely in the cerebral cortex, hypothalamus, thalamus, and HC on both somata and dendrites (Neumaier et al., 2001). Gustafson et al. (1996) report that 5-HT(7)Rs are densely located in the rat limbic system but also occur elsewhere in the thalamus, hypothalamus, and HC. This receptor is the predominant one in rat astrocytes, where it is coupled with adenylate cyclase (Hirst et al., 1997). 5-HT(6)Rs are located (þþ) in rat corpus striatum, nucleus accumbens, and olfactory tubercle; (þ) in certain areas of the cerebral cortex, the HC, thalamus and SN, and hypothalamus; and at very low concentrations in the cerebellum, midbrain, and globus pallidus (Roberts et al., 2002). Gerard et al. (1997) found 5-HT(6)Rs particularly in rat brain extrapyramidal and limbic areas. In the CNS, 5-HT(3)Rs are mostly presynaptic on axon terminal and fibers of nonmonoaminergic neurons (Kidd et al., 1993). In the rat telencephalon and cerebral cortex, 5-HT(3)Rs are located mainly on GABAergic INs that contain CCK (but not SS) and calbindin (but not parvalbumin) (Morales and Bloom, 1997). This receptor is the only 5-HT receptor that controls an ion channel. At presynaptic sites, it induces Ca2þ influx and is permeable to Ca2þ ions, whereas at postsynaptic loci, it induces depolarization and is permeable to Naþ and Kþ ions (Ronde and Nichols, 1998; Sodhi and Sanders-Bush, 2004). In HC CA1 stratum radiatum, 5-HT directly excites INs by means of 5-HT(3)Rs, especially when the cells are depolarized (McMahon and Kauer, 1993). In most cases, inhibitory autoreceptors on an axon terminal are thought to mediate negative feedback inhibition of release of that transmitter. However, Kalsner and Abdali (2002) present evidence that in the rabbit, somatodendritic sites on raphe 5-HT neurons bear inhibitory receptors for 5-HT release that operate independently from such feedback.

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D. Synaptic Versus Volume Transmission In diVerent loci, 5-HT terminals can either form synapses or operate nonsynaptically by volume transmission. For example, in the mouse suprachiasmatic nucleus, most 5-HT(1B)Rs and 5-HT(7)Rs are extrasynaptic (Belenky and Pickard, 2001). In rat periacqueductal gray (PAG), the 5-HT axons to the output neurons operate by means of volume transmission (Lovick et al., 2000). This is also the case in rat HC CA3 and dentate gyrus (Oleskevich et al., 1991) and in rat CA1 stratum radiatum in contrast to GABAergic terminals, which are always synaptic (Umbriaco et al., 1995). In monkey PFC, only 23% of the abundant 5-HT axons form synapses (asymmetrical mainly on the dendritic shafts of INs). Most operate by volume transmission (Smiley and GoldmanRakic, 1996). In contrast again, DA axons in this location form synapses mostly onto the dendrites of pyramidal cells. Jansson et al. (2001) also report that in rat forebrain, 5-HT(2A)R–mediated activity is by volume transmission. Some extrasynaptic 5-HT transmission is also reported in rat HC, raphe, and septum (Kia et al., 1996) and for 5-HT(1A)Rs in the rat raphe (Riad et al., 2000). In contrast, the 5-HT axons from the raphe running to the SNpc form only asymmetrical synapses located on the spines and dendritic shafts of DA neurons with no volume transmission (Corvaja et al., 1993). Likewise, in rat lateral septum, most (95%) 5-HT varicosities make symmetrical synapses on neuronal somata, dendritic shafts, and spines with no volume transmission (Dinopoullos et al., 1993).

II. Electrical Actions

A. Potassium Related 5-HT, through its various receptors, exerts complex eVects on the electrical activity of neurons. Newberry et al. (1999), using human neocortical slices, found that the application of 5-HT produced a variety of eVects on 21 individual neurons. 1. It depolarized 11 of 21 neurons by inhibiting the resting Kþ conductance by means of 5-HT(2A)Rs. 2. It hyperpolarized 5 of 21 neurons by opening ion channels by means of 5-HT(1A)Rs. This also occurred in rat SCN (Jiang et al., 2000). 3. It had a biphasic eVect on two neurons. 4. It had no eVect on the remaining three neurons. Likewise, in rat somatosensory and motor cortex, 5-HT depolarized some neurons and hyperpolarized others (Foehring et al., 2002). Craven et al. (2001)

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confirmed in guinea-pig raphe neurons that activation of 5-HT(1A)Rs hyperpolarized the cell by means of a G-protein mediated inwardly rectifying potassium (GIRK) inhibitory current. This is eVected by direct PKA-mediated phosphorylation of the channel (Wischmeyer and Karschin, 1996). This also occurs in rat dorsolateral septal nucleus (Goto, 1998), in rat ventrolateral medullary nucleus (Hwang and Dun, 1999), in rat neocortex (Takigawa and Alzheimer, 1999), and in rat HC CA1 neurons (Gasparini and Di Francesco, 1999). Bobker and Williams (1995) found that 5-HT inhibition in guinea-pig nucleus prepositus hypoglossi is mediated by a rapidly activated GIRK current and by a slowly activated outwardly rectifying Kþ current, both of which cause IPSPs. 5-HT–mediated inhibition in ferret thalamus is mediated directly by promoting Kþ conductance by means of activation of the 5-HT(1A)R and also indirectly by activation of inhibitory GABAergic INs (Monckton and McCormick, 2002). In guinea pig spinal trigeminal nucleus, 5-HT (and NE) inhibit 70% of neurons by a hyperpolarization mediated by an increase in a Kþ conductance. 5-HT and NE also activate GABAergic (in 20% of recordings) and glycinergic INs (in 20% of recordings) to cause IPSPs in separate groups of cells (Grudt et al., 1995). Activation of 5-HT(2)Rs in guinea-pig V motoneurons depolarized the cell by inhibiting a resting Kþ leak conductance partly by means of 5-HT(2)Rs (Hsiao et al., 1997). Other 5-HT receptors have various eVects. The 5-HT(4)R increases membrane excitability in the rat hippocampus by inhibiting the Caþþ-activated Kþ current responsible for slow afterhyperpolarization by means of increasing cAMP levels and activating PKA (Chapin et al., 2002; Siarey et al., 1995; Torres et al., 1995) and without involvement of I(h) unlike many other neurons (Chapin et al., 2002). In rat motoneurons, 5-HT inhibits medium duration afterhyperpolarization by means of cAMP-dependent activation of PKC (Inoue et al., 1999). In the rat HC CA3, 5-HT(7)Rs directly inhibit a Ca2þ activated Kþ channel, resulting in the modulation of synchronized network activity in the hippocampus (Gill et al., 2002). 5-HT(2C)s inhibit an inwardly rectifying Kþ current expressed in Xenopus oocytes by means of PKC (DiMagno et al., 1996). 5-HT, as well as adenosine, somatostatin, GABA(B)Rs, and 50% of mGluRs, all activate inwardly rectifying Kþ conductance acting synergistically (Sodickson and Bean, 1998). In rat dorsal raphe, 5-HT induces a GIRK current without involvement of Ca2þ or PKC (Katayama et al., 1997). In rat C6 glioma cells, 5-HT(2A)Rs increase i.c. phosphoinositol hydrolysis and increase i.c. Ca2þ levels, leading to activation of an outward Kþ current (Elliott et al., 1995). Most 5-HT neurons in rat raphe contain two new types of two pore domain Kþ channels, called TaSK-1 and TASK-3, with unique properties. They generate instantaneous open-rectifier (‘‘leak’’) currents that are modulated by external pH (Washburn et al., 2002).

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I(h) is the hyperpolarization-activated nonselective cation current that depolarizes the membrane. It acts as a pacemaker current in neuronal membranes (Funahashi et al., 2003; Pal et al., 2003). Transient deactivation of I(h) induces afterhyperpolarization (AHP). GIRK channel activation inhibits I(h)-dependent AHP (Takigawa and Alzheimer, 2003). B. Role of Calcium Channels 1. 5-HT(1) Receptors 5-HT also modulates neuronal activity by its action on Ca2þ channels that aVect oscillatory activity and neuronal firing. The 5-HT(1A)R inhibits N- and P-type Ca2þ channels by means of a G-protein and so reduces the amplitude of the afterhyperpolarization (AHP) in rat motoneurons inter alia (Bayliss et al., 1995). In rat ventromedial thalamus (Rhee et al., 1996) and rat melanotrophs (Ciranna et al., 1996), 5-HT(1A)Rs also inhibit N- and Q-type Ca2þ channels by means of a PTX-sensitive G-protein. In neocortical pyramidal cells, 5-HT inhibits N- and P(but not L-) type Ca2þ currents (Foehring, 1996). In rat caudal raphe, 5-HT inhibits N- and P/Q-type Ca2þ channels and AHPS (Bayliss et al., 1997). In general, activation of 5-HT(1A)Rs attenuates the activity of both N-type Ca2þ channels and NMDARs, thus lending protection against excitotoxicity. However, this is not the only protective mechanism involved. In a Ca2þ channel–deficient striatal cell line (HN2-5), agonist binding to 5-HT(1A)Rs results in a MAPKmediated inhibition of a caspase 3–like enzyme and a 60 to 70% suppression of anoxia-induced apoptosis by means of a Ca2þ channel–independent mechanism (Adayev et al., 1999). In guinea pig nucleus of Meynert neurons, 5-HT(1A)Rs block N-type Ca2þ currents by means of G(i) and G(o) subtypes of G-protein (Williams et al., 1998). In rat HC CA3 cells, presynaptic 5-HT(1A)Rs inhibit Ca2þ entry and so diminish glutamatergic synaptic transmission and its associated EPSPs (Schmitz et al., 1995). In rat raphe, activation of PKC inhibits the 5-HT(1A)R–induced Ca2þ influx but does not aVect the Kþ current (Chen and Penington, 1996). In guinea pig, NOSþ ACh neurons, 5-HT depresses the increase in i.c. calcium in somata and dendrites produced by action potentials. It does this both directly by inhibiting spike-evoked Ca2þ transients and indirectly by reducing firing rates (Leonard et al., 2000). In mouse glioma NG10-15 cells, Jones and Yakel (1998) demonstrated cross-talk between Ca2þ levels and 5-HT3Rs. Ca2þ influx by means of a VSCC increases i.c. Ca2þ levels. This leads to a dramatic slowing in the kinetics of desensitization of 5-HT(3)Rs. In rat astrocytes from the cerebral cortex, 5-HT evoked a spike in i.c. Ca2þ levels followed by a lower plateau. The former was caused by Ca2þ release from

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internal stores and the latter by Ca2þ entry by means of voltage-independent Ca2þ channels (Hagberg et al., 1998). 2. Other 5-HT Receptors In rat neocortical layer V pyramidal cells, 5-HT(2A)Rs induce EPSPs in apical dendrites by both presynaptic and postsynaptic mechanisms (Aghajanian and Marek, 1997). In rat PFC layer V and VI pyramidal neurons, 5-HT(2)Rs inhibit L-type Ca2þ currents by means of the cascade: q G-protein ! PLC ! phosphoinositol synthesis ! mobilization of Ca2þ from intracellular stores and activation of calcineurin (Day et al., 2002). In rat amygdala, 5-HT(1A)Rs inhibit voltage-sensitive Ca2þ channels (VSCCs) (Lin et al., 2001). In pyramidal cells of layer V of the rat frontal cortex, 5-HT excites output neurons by raising the frequency and amplitude of EPSCs by means of 5-HT(2A)Rs (Lambe et al., 2000). Zhou and Hablitz (1999) confirm that 5-HT(2A)Rs strongly promote spontaneous EPSPs (as well as spontaneous IPSPs) in cortical pyramidal cells but only weakly in the case of INs. However, 5-HT only weakly inhibited evoked EPSPs and IPSPs. In these experiments, 5-HT(3)Rs activated a subset of INs. In contrast, in rat ventrolateral medulla, 5-HT(1B and 1D)Rs inhibit EPSPs in most cells and promote IPSPs in others (Huang and Dun, 1999). In the superficial layers of the rat medial entorhinal cortex, 5-HT (mainly by means of 1ARs) strongly inhibits EPSPs by a presynaptic mechanism (Schmitz et al., 1999). 5-HT (1A)Rs tonically inhibit the spontaneous firing of HC CA1 pyramidal cells in quiet awake rats (Kasamo et al., 2001). In mouse PFC pyramidal cells, activation of 5-HT(2A and 2C)Rs reduced rapidly inactivating Naþ current by lowering maximum current amplitude and shifting fast inactivation voltage dependence. This eVect was mediated by G(q) ! PLC ! PKC. The activation of 5-HT(2A/ C)Rs did not aVect somatic action potentials of these neurons, but it had the important eVect of reducing the amplitude of back-propagated actin potentials into the dendritic, the significance of which will be shown later (Carr et al., 2002). In rat HC CA1 pyramidal neurons, 5-HT(7) and 5-HT(1A) Rs promote the hyperpolarization-activated nonselective cation current called I(h), but 5-HT(4) Rs decrease it (Bickmeyer et al., 2002). In rat CA1 cells, 5-HT(4)Rs raise cell excitability by inhibiting slow Ca2þ-activated AHPs (by means of cAMP and PKA) This is mediated by inhibiting the ability of extracellular Ca2þ to trigger Ca2þ release from i.c. stores (Torres et al., 1996). In rat striatal synaptosomes, activation of 5-HT(3)Rs raises i.c. Ca2þ levels slowly (Nichols and Mollard, 1996). In rat HC CA3 pyramidal neurons, activation of 5-HT(7)Rs inhibits the slow AHP induced by Ca2þ spikes (Bacon and Beck, 2000). 5-HT(7)Rs depolarize thalamic anterodorsal neurons by raising I(h) by means of a cAMP-dependent and PKA-independent mechanism (Chapin and Andrade, 2001a,b). In the LGN, 5-HT upregulates the hyperpolarization-activated current I(h) in a Ca2þ-dependent manner. In the anterior thalamus, this same eVect is mediated

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in a Ca2þ-independent manner (Chapin and Andrade, 2000). In rat astrocytes, activation of 5-HT(2B)Rs increases i.c. Ca2þ levels from internal stores (Sanden et al., 2000). In the rat tuberomammillary hypothalamic nucleus, 5-HT(2C)Rs excite neurons by activating an electrogenic Naþ/Ca2þ exchanger that results in a net inward current (Eriksson et al., 2001). 5-HT(2)Rs inhibit the hyperpolarization-activated current (Ih) by means of PKC (Liu et al., 2003). In rat HC astroglial gap junctions, the velocity of induced Ca2þ waves is increased by 5-HT and glu (Blomstrand et al., 1999). In rat raphe, 5-HT neurons need some input to fire. This can be provided by 1 adrenergic receptors, histamine H1Rs, and oxytocin/hypocretin OX2Rs (Brown et al., 2002). In rat hypoglossal nucleus motoneurons, activation of 5-HT1A receptors primarily reduced voltage-activated Ca2þ signals without a significant impact on basal [Ca]i. In contrast, activation of 5-HT2 receptors initiated a net inward current followed by membrane depolarization (Ladewig et al., 2004). 3. Serotonin and Neuronal Oscillation Serotonin can also modulate oscillatory behavior in neurons. For example, in rat inferior olive, 5-HT(2)Rs inhibit oscillations by depressing both T-type Ca2þ currents and a resting Kþ current (fast IK/ir) and by promoting the cation current I(h) (Placantonakis et al., 2000). Serotonin modulates the 200 Hz ‘‘ripple’’ oscillations in behaving rat HC CA1. A 5-HT(1A)R antagonist reduced the occurrence and the intrinsic frequency of ripples, whereas a 5-HT(3)R antagonist increased the number but reduced the frequency of the ripples (Ponomarenko et al., 2003). 4. 5-HT Release Okada et al. (2001) showed in microdialysis studies in rat HC that basal and Kþ-evoked 5-HT release can be modified by two pathways (1) (major) N-type voltage-sensitive Ca2þ channels (VSCCs) ! phospholipid-dependent activation of PKC/syntaxin and (2) (minor) P-type VSCC ! cAMP-dependent PKA/ synaptobrevin. In addition, they found that activation of adenosine A1-Rs inhibited 5-HT release by both pathways, whereas activation of A2-Rs promoted 5-HT release only by pathway (2). Bagdy and Harsing (1995) showed that in the rat raphe, dendritic release of 5-HT is mediated by 5-HT(1A)Rs and N-type VSCCs, whereas in the HC, the release of 5-HT from axon terminals is mediated by 5-HT(1B)Rs. They also suggested that the 5-HT(1A)R–mediated inhibition of 5-HT release may be due to the opening of VSCCs. 5-HT release in rat dorsal raphe depends on opening fast-activated Naþ channels (Matos et al., 1996). 5. Miscellaneous Electrical EVects of 5-HT Serotonin has various eVects on the electrical behavior of neurons depending on circumstances. In the rat, serotonin activates most neurons in the subthalamic nucleus (Flores et al., 1995).

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III. Interactions Between Serotonin and Other Neurotransmitter Systems

A. Actions of Serotonin on Glutamate Synaptic Transmission Serotonin usually inhibits glutamatergic synaptic transmission, either presynaptically or postsynaptically, but it can also promote such transmission in certain loci. 1. Basic Anatomy In rat medial nucleus of the solitary tract (NTS), many neurons carry both NMDARs and 5-HT(2A)Rs but in diVerent locations. The former are only postsynaptic, whereas some of the later are extrasynaptic (Huang and Pickel, 2002). Interactions between these two receptors modulate the excitability of NTS neurons both at the presynaptic and postsynaptic levels (Huang and Pickel, 2003). Single 5-HT neurons in culture can release both 5-HT and glu at the synapses they form with other neurons in the culture and may do so in vivo ( Johnson and Yee, 1995). In rat cerebral cortex, 5-HT modulates plasticity by regulating the density of AMPAR subtypes (R1 up; R2/3 down) (Shutoh et al., 2000). 5-HT(6) and 5-HT(7)Rs are located in limbic areas where mGluRs modulate their mRNA expression (Healy and Meador-WoodruV, 1999). 2. Presynaptic Inhibition In rat HC, stimulation of 5-HT(1A)Rs inhibits evoked glu release and so protects the cell against excitotoxic neurotoxicity (Mauler et al., 2001). In rat raphe, the 5-HT and glu systems have a complex relationship. Local glu neurons excite 5-HT neurons by means of AMPARs. A retrocollateral of the 5-HT cell axons inhibits (by means of 5-HT[1B]Rs) the incoming glu fiber in a negative feedback loop (Li and Bayliss, 1998). Likewise, in the medium spiny neurons of the rat nucleus accumbens (N.Ac.), 5-HT(1B)Rs inhibit electrically evoked (glu-mediated) EPSCs by a presynaptic mechanism (Muramatsu et al., 1998). In rat XII motoneurons, 5-HT(1R)Rs inhibit glu synaptic transmission by decreasing the probability of glu release (Singer et al., 1996). In rat cerebellum and human cerebral cortex, 5-HT(1D)Rs presynaptically inhibit glu release (Maura and Raiteri, 1996; Maura et al., 1998). In rat subicular cortex, 5-HT(1B)Rs inhibit low-, but not high-, frequency synaptic transmission by inhibiting glu release from glu axons derived from CAI pyramidal cells (Boeijinga et al., 1996). In neonatal mouse sensory cortex layer IV, 5-HT(1B)Rs inhibit glu release presynaptically. This could favor the propagation of high-frequency thalamocortical activity (Laurent et al., 2002). In the mouse, 5-HT(1B)Rs function both as autoreceptors and heteroreceptors. Activation of this receptor in the dorsal raphe potently inhibits the slow IPSPs evoked by 5-HT(1A)Rs. It also inhibits glu release in the nucleus accumbens and inhibits GABA release in the ventral

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midbrain (Morikawa et al., 2000). In rat locus ceruleus (LC), 5-HT axon terminals directly contact LC neurons. 5-HT inhibits glu excitation of these NE cells by both presynaptic and postsynaptic mechanisms, as well as by indirect actions by means of glia (van Bockstaele, 2000). In the SCN, 5-HT inhibits glu release, but higher doses can raise i.c. Ca2þ levels in some SCN neurons by means of modulation of GABA cell activity (Flett and Colwell, 1999). In layers II and III of rat entorhinal cortex, 5-HT(1A)Rs powerfully inhibit, by a presynaptic mechanism, synaptic transmission as mediated by both AMPARs and NMDARs (Schmitz et al., 1998). In rat basolateral amygdala, 5-HT(1A)Rs on excitatory glu terminals inhibit glu release by means of cAMP-dependent mechanisms and modulation of VOCCs (Cheng et al., 1998). 3. Presynaptic Facilitation In the rat HC-septal pathway, 5-HT(1A)Rs on lateral septal nucleus neurons enhance the eYciency of glu transmission (Hasuo and Akasu, 2001). Activation of 5-HT(2A)Rs in this locus promotes the release of glu by means of links with PKC and thus induces EPSPs (Hasuo et al., 2002). In rat frontal cortex layer V pyramidal cells, 5-HT powerfully induces glu release from thalamocortical terminals by blocking Kvl.2 Kþ channels (Lambe and Aghajanian, 2001; Lambe et al., 2000). In rat medial PFC layer V, 5-HT(2A) Rs activate pyramidal cells by a presynaptic mechanism that increases glu release onto apical dendrites. This eVect is inhibited by adenosine (Stutzmann et al., 2001). 4. Postsynaptic Inhibition In rat visual cortex layer V pyramidal cells, 5-HT inhibits the EPSCs induced by glu by reducing their frequency but not their amplitude (Murakoshi et al., 2001). In rat PFC pyramidal cells, 5-HT(1A)Rs inhibit AMPA-evoked currents by means of downregulation of Ca2þ/calmodulin–dependent PKII. This action inhibits the phosphorylation of the AMPA subunit gluR1 in a protein phosphatase-I–dependent manner (Cai et al., 2002b). In rat neocortical cultures, 5-HT (1A)Rs inhibit (and so protect against) glu neurotoxicity and downregulate NMDA-induced cGMP accumulation (Gandolfi et al., 2002). In rat cerebellum and human cortex, 5-HT(1A)Rs inhibit the cGMP response to stimulation of AMPARs and NMDARs (Maura and Raiteri, 1996; Maura et al., 1998). In rat cerebral cortex slices, 5-HT(1B/1D)Rs inhibit the NMDA ! NO ! cGMP cascade (Stepien et al., 1999). In rat cerebellum, this cascade is inhibited by 5-HT (receptor unspecified) (Maura et al., 1995) and by 5-HT(2C)Rs (Marcoli et al., 1997). In rat lateral entorhinal cortex, 5-HT(1A)Rs hyperpolarize and thus inhibit pyramidal cells both by modulating Kþ conductance and by inhibiting glu transmission (Grunschlag et al., 1997). In rat red nucleus, 5-HT(1A)Rs inhibit glu-evoked excitation in 85% of neurons acting mainly on non-NMDARs. The eVect here of 5-HT on NMDAR-evoked activity varies in diVerent cells

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(excitation, inhibition, or no action) (Licata et al., 1998). In rat basolateral amygdala, 5-HT inhibits glu-evoked action potentials, but in this locus it does so only in the presence of corticosteroids (Stutzmann et al., 1998). In guinea pig dentate gyrus slices, release of glu from the slice is inhibited by postsynaptic 5-HT (1A)Rs and is promoted by NMDARs (Matsuyama et al., 1996). In rat spinal cord dorsal horn, 5-HT inhibits glu-induced EPSCs by means of a G-protein. This diminishes the pain response (Li and Zhuo, 2001). In mouse dorsal raphe, 5-HT (1B)Rs inhibit the slow IPSPs produced in this locus by stimulation of 5-HT(1A) Rs (Morikawa et al., 2000). In rat isolated SCN neurons, 5-HT(7)Rs suppress gluinduced responses as measured by increased i.c. Ca2þ levels (Quintero and MacMahon, 1999). Jiang et al. (2000) report that 5-HT regulates the activity of SCN neurons by inhibitory presynaptic and inhibitory postsynaptic activity. In the SCN, the activity of 5-HT(1B) autoreceptors exhibit a circadian rhythm. Serotonin release in the SCN, evoked by stimulation of the raphe nucleus, attenuates the increase in FOS expression in the SCN induced by light (MeyerBernstein and Morin, 1999). Prosser (2001) reviews the complex system by which glutamate blocks the serotoninergic phase advances of the mammalian circadian pacemaker through interacting AMPA and NMDA receptors. 5. Postsynaptic Facilitation The only reported instance, to my knowledge, is quoted previously where the eVect here of 5-HT on NMDAR-evoked activity varies in diVerent cells including excitation on some cells (Licata et al., 1998). 6. Unspecified Inhibition In rat cortical slices, 5-HT(1A)Rs inhibit corticostriatal neurons (Dijk et al., 1995). In rat striatum, 5-HT has a tonic inhibitory action on glu transmission but has no eVect on DA transmission (Di Cara et al., 2001). Throughout the cerebellar nuclei in the cat brain, a dense plexus of 5-HT axons inhibits NMDA-evoked currents and promotes GABA-evoked currents (Kitzman and Bishop, 1997). 7. Unspecified Facilitation In rat subiculum, 5-HT excites 5-HT(2B)Rs on glu neurons that project to the nucleus accumbens, where they synapse on DA terminals by NMDARs and here promote DA release (Boulenguez et al., 1998). B. Actions of Glutamate on Serotonin Systems Glutamate has mainly inhibitory actions on serotonin systems. In freely moving rats using microdialysis, Smith and Whitton (2000) showed that a low dose of NMDA in the raphe inhibited 5-HT release, but a high dose promoted

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such release. In the cerebral cortex, NMDA had the opposite eVect (low doses promoted and high doses inhibited 5-HT release). In freely moving rats, activation of NMDARs in the HC inhibited 5-HT release by 70% by means of NO in ways described by the authors as ‘‘complex’’ (Segieth et al., 2001). In rat periaqueductal gray, NMDARS inhibit, whereas mGluRs increase, 5-HT release. This eVect is probably eVected by means of an intermediate GABA interneuron (IN ) (Maione et al., 1998). In rat medial PFC layer V pyramidal cells, activation of mGlu2/3Rs inhibits the EPSPs induced by the presynaptic release of glu induced by stimulation of 5-HT(2A)Rs. The authors comment on the ‘‘striking overlap’’ of mGlu2/3Rs and 5-HT(2A)Rs in the medial PFC that is not apparent in other cortical areas (Marek et al., 2000). GluRepsilon1 mutant mice have malfunctioning NMDARs and overactive 5-HT (and DA) systems in the cerebral cortex and striatum (Miyamoto et al., 2001). In astrocytes in culture, activation of mGluRs inhibit 5-HT–evoked rises in i.c. Ca2þ levels (Haak et al., 1997). A review of the role of 5-HT in astrocyte hyperpolarization is given by Laming et al. (2000). In contrast, in rat cortical slices, stimulation of NMDARs and non-NMDARs promotes 5-HT release partly from serotoninergic nerve terminals (Fink et al., 1995). In rat striatum, stimulation of AMPARs (as well as of 5-HT(2A/C)Rs) promotes 5-HT release (Abellan et al., 2000). Neurons in the rat nucleus accumbens receive inhibitory 5-HT input (onto 5-HT[1A]Rs), as well as inhibitory DA input and excitatory glu input from the parafascicular thalamic nucleus (Nagaoka et al., 1998).

C. Actions of Serotonin on GABA Systems 1. Presynaptic Inhibitory In guinea pig dentate gyrus, 5-HT(1A)Rs inhibit GABA release (Matsuyama et al., 1997), and in rat VTA, 5-HT(1B)Rs also inhibit GABA release (Yan and Yan, 2001). In rat basolateral amygdala, presynaptic 5-HT(1A)Rs inhibit GABAergic miniature IPSCs by inactivating the cAMP pathway. This acts directly on the GABA-releasing process independently of Kþ and Ca2þ channels (Koyama et al., 1999). These receptors, by means of a G-protein–mediated mechanism, also inhibit the transient increase in miniature IPSCs produced in this locus by stimulation of 5-HT(3)Rs (Koyama et al., 2002). 2. Presynaptic Excitatory In rat frontal cortex slices, 5-HT(2A)Rs promote GABA release (Cozzi and Nichols, 1996). In rat amygdala 5-HT(3)Rs increase GABA release by a presynaptic (and no postsynaptic) action. This involves the influx of Ca2þ through the 5-HT(3)R–associated ionic channel (Koyama et al., 2000).

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3. Postsynaptic Inhibitory The ventromedial PFC exerts a powerful inhibitory action on 5-HT neurons in the dorsal raphe by releasing 5-HT, which activates inhibitory 5-HT(1A)Rs (Celada et al., 2001), and by excitation of GABAergic INs (Celada et al., 2001; Varga et al., 2001). In rat nucleus accumbens shell, 5-HT postsynaptically inhibits GABAergic neurons (Van Bockstaele et al., 1996). 4. Postsynaptic Excitatory In rat medial septum/nucleus of the diagonal band of Broca (NDBB), 5-HT activates GABAergic neurons directly using multiple types of 5-HTRs (Alreja, 1996). In rat lateral amygdala, 5-HT activates GABAergic INs that inhibit the glu input from the auditory thalamus and cerebral cortex. This action is related to the control of fear conditioning (Stutzmann and LeDoux, 1999). In the PFC pyramidal cells, 5-HT(2)Rs inhibit GABAergic currents by phosphorylation of GABARs by means of activation of PKC anchored to the scaVolding protein RACK (receptor for activated C kinase) (Feng et al., 2001). In rat neocortex, 5-HT(3A)Rs are found exclusively on a subset of GABA INs that coexpress VIP/ CCK. 5-HT here induces a fast membrane depolarization by activating an inward current, possibly by means of synaptic contacts. These neurons are also excited by nicotinic agonists, indicating a convergence on these cells of fast 5-HT and cholinergic transmissions (Ferezou et al., 2002). In the rat raphe-HC projection system, 5-HT(3)Rs directly excite HC GABAergic INs in the stratum radiatum of CA1. These neurons project widely to the HC and dentate gyrus (McMahon and Kauer, 1997). 5. Presynaptic Both Excitation and Inhibition In guinea pig HC slices, 5-HT(4)Rs have complex eVects on GABA release. They have no eVect on basal release, but on electrically stimulated release, low concentrations of 5-HT promote GABA release (indirectly by means of M1 and M3 AchRs), whereas high concentrations inhibit GABA release (indirectly by means of M2 AChRs) (Bianchi et al., 2002). 6. Postsynaptic Both Excitation and Inhibition In rat PFC pyramidal cells, 5-HT(4)Rs regulate PKA-induced phosphorylation of GABA(A)Rs. The direction of modulation (up/down) depends on the basal level of PKA activation; elevation of PKA activity changes enhancement to reduction (Cai et al., 2002a). Activation of raphe 5-HT neurons in rats desynchronize HC . Two pathways mediate this activity (1) inhibitory by means of 5-HT(2A)R activation of inhibitory GABAergic INs in the HC (exclusively on their apical dendrites) leading to the inhibition of HC neurons and (2) disinhibition by 5-HT activation of GABAergic INs in the medial septum/NDBB, which

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in turn inhibit inhibitory HC basket and chandelier cells that normally inhibit HC pyramidal cells. All this results in the activation of the latter cells (Leranth and Vertes, 1999). 7. Unspecified Inhibition In rat HC CA1 pyramidal cells, the activating eVect of the 5-HT antagonist zacopride depends on 5-HT(3)Rs–mediated down regulation of GABAergic inhibition plus 5-HT(4)R–mediated reduction of Ca2þ-mediated Kþ conductance, which combine to increase pyramidal cell excitability (Siarey et al., 1995). 8. Unspecified Excitation In cat cerebellum, 5-HT potentiates GABA-evoked currents (Kitzman and Bishop, 1997). 9. Miscellaneous Items In primate cerebral cortex, GABAergic chandelier neurons make inhibitory synaptic contacts, whereas 5-HT(1A)Rs make inhibitory paracrine contacts (DeFelipe et al., 2001). In rat HC, 5-HT agonists regulate GABA transporter (GAT1) function by means of PKC (Beckman et al., 1999). In rat nucleus accumbens shell, 36% of 5-HT terminals converge on a common dendrite together with a GABA terminal, suggesting that these two terminals may interact at the presynaptic level (Van Bockstaele et al., 1996). In rat dorsal raphe, neurons expressing GABA(B) receptor RNA are very abundant, and 855 of these coexpress 5-HT transporter RNA (Serrats et al., 2003). D. Actions of GABA on Serotonin Systems In all locations, GABA strongly inhibits 5-HT neurons both at the presynaptic and postsynaptic level. GABA(B)Rs inhibit 5-HT neurons in rat raphe and so promote HC (Varga et al., 2002). In cat raphe, GABAergic neurons inhibit 5-HT neurons during REM sleep (Torterolo et al., 2000). In rat dorsal raphe and nucleus accumbens, GABA agonists strongly inhibit 5-HT release (Tao and Auerbach, 2000). E. Actions of Serotonin on the Cholinergic System 1. Presynaptic In rat cerebral cortex, 5-HT(3)Rs inhibit ACh release (but not 5-HT nor DA release) (Crespi et al., 1997). In the frontoparietal cortex and HC of freely moving rats, activation of 5-HT(3)Rs had no eVect on spontaneous ACh release but

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diminished Kþ-activated ACh release (Giovanni et al., 1998). Likewise, in guinea pig brain slices, activation of ACh(4)Rs does not aVect resting ACh outflow but increases the outflow produced by electrical stimulation (Siniscalchi et al., 1999). In guinea pig SN, 5-HT released from raphe-nigral aVerents promotes the release of AChE from the dendrites of DA neurons (Dickie and Greenfield, 1994). 2. Postsynaptic In rat medial septum and NDBB, 5-HT suppresses slow afterhyperpolarization in cholinergic neurons and alters the firing pattern from a slow regular discharge to one exhibiting a modest spike frequency adaptation (Gorelova and Reiner, 1996). In rat HC slices, 5-HT(1A)Rs inhibit muscarinic-induced increases in PI turnover by means of the PLA2 ! AA ! PKCgamma cascade (Claustre et al., 1991). In the rat, 5-HT neurons in the dorsal raphe and ACh neurons in the LDTN anatomically interconnect and may reciprocally influence each other by means of synaptic connectivity (Wang et al., 2000). 3. Complex Interactions In rat striatal slices, 5-HT(2A)Rs increase D1-activated ACh release by means of interposed collaterals of substance P–containing neurons that raise ACh eZux by activating neurokinin receptors on cholinergic INs (Ramirez et al., 1997). F. Actions of Serotonin on the Dopamine System Serotonin receptors usually promote but can decrease dopamine release, depending on circumstances. In rat olfactory tubercle and striatum, 5-HT(3)Rs upregulate both basal and Kþ-evoked DA release (Zazpe et al., 1994). Likewise, in rat striatum, 5-HT promotes DA release partly by action on the DA reuptake site (De Deurwaedere et al., 1996). In rats, inhibitors of 5-HT uptake enhance DA eZux in N.Ac. and increase cocaine-induced hyperactivity (Bubar et al., 2003). In rat medial PFC and N.Ac., 5-HT(2A)Rs increase amphetamineinduced DA release (Kuroki et al., 2003). In the posterior (but not the anterior) nucleus accumbens of the rat, 5-HT(2)Rs increase DA release (Bowers et al., 2000). In rat SN and striatum, 5-HT promotes DA release from dendrites by a mechanism that is uncoupled from the nerve impulse–dependent control of nerve terminal DA release (Codd and Abercrombie, 2003). In rat striatum, Lucas et al. (2000) found that 5-HT does not aVect basal DA release but upregulates DA outflow produced by stimulation of the nigrostriatal pathway. In mouse striatum, 5-HT both promotes and inhibits DA release by diVerent mechanisms. Inhibitory presynaptic 5-HT heteroreceptors and both inhibitory and stimulatory somatodendritic 5-HT receptors regulate DA release. Furthermore,

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5-HT also promotes DA release by reversing the DAT. 5-HT(2)Rs are mainly involved in this eVect (Sershen et al., 2000). 5-HT(3)Rs can colocalize with nicotinic receptors on presynaptic DA terminals in the striatum and modulate DA release (Nayak et al., 2000). In rat VTA, 44% of 5-HT(2A)Rs are located on dendrites and 56% on axons of DA cells (Doherty and Pickel, 2000). In rat raphe, 5-HT(1A)Rs inhibit DA-related behaviors such as the acoustic startle reflex (Meloni and Davis, 2000). In the dorsal striatum, 5-HT(1B)Rs are not involved in DA release in this brain area in mice (De Groote et al., 2003). In rat N.Ac., 5-HT(3)Rs inhibit DA release (Campbell and McBride, 1995). G. Actions of Serotonin on the Norepinephrine System In rat HC, 5-HT(3)Rs inhibit NE release (Matsumoto et al., 1995). In rat locus ceruleus, 5-HT(1A)Rs activate NE neurons (Hamamura et al., 1997). This involves the following complex mechanism (Szabo and Blier, 2001a, b). Serotonin activates inhibitory 5-HT(1A)Rs on glutamate neurons, thereby lowering glu release and depressing the activity of glu receptors (possibly of the kainate type) on 5-HT terminals. The ensuing decrease in 5-HT release then lowers the activity of 5-HT(2)Rs on GABA neurons that project to NE neurons, thus leading to a disinhibition of the latter (Szabo and Blier, 2001a, b). In rat locus coeruleus, 5-HT(1A)Rs activate NE neurons (as measured by increased Fos-like immunoreactivity) and so increase NE transmission in the forebrain, especially in the medial PFC (Hajos-Korcsok and Sharp, 1999). In general, 5-HT(1B) agonists raise Fos expression in the rat striatum (Wirtshafter and Cook, 1998). Microdialysis of the HC in living rats by 5-HT stimulates 5-HT-3Rs and so inhibits NE release (Matsumoto et al., 1995). H. Effects of Serotonin on the Nitric Oxide System This interaction was inhibitory in three instances reported. In C6 glioma cells, stimulation of 5-HT(2A)Rs inhibited NOS activity induced by cytokines (Miller and Gonzalez, 1998) and by LPS (by means of PKC) (Miller et al., 1997). In rat HC synaptosomes, 5-HT(1A)Rs strongly inhibited NO/cGMP synthesis (as well as NMDA-evoked AA release; Strosznajder et al., 1996). I. Effects of Nitric Oxide on the Serotonin System This eVect varies in diVerent systems. In rat synaptosomes, NO inhibits 5-HT uptake (Asano et al., 1997). In freely moving rats, NO lowers levels of 5-HT and of DA (but not by means of a cGMP-dependent mechanism) (Wegener et al., 2000).

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In rat locus ceruleus, NO promotes the release of 5-HT but only if applied concomitantly with EAA activity or stress (Sinner et al., 2001). In rat hypothalamus, endogenous NO modulates the release of 5-HT in a biphasic manner (low dose inhibits; high dose promotes) in a cGMP-dependant mechanism (Kaehler et al., 1999). Smith and Whitton (2000), working with freely moving rats, reported that NO modulates NMDA-evoked 5-HT release in a complex manner. In the raphe, low doses of NMDA decrease 5-HT release, whereas high doses increase it. However, in the cerebral cortex, the converse eVect was observed. Coinfusion of the NOS inhibitor 7-NI abolished the eVect of high, but not low, doses of NMDA. 7-NI given alone decreased 5-HT release in the raphe but increased it in the frontal cortex. Low doses of the NO donor SNAP infused into the raphe decreased 5-HT release locally but increased it in the frontal cortex. High doses of SNAP had the opposite eVect in both brain regions. Similar results in the HC of freely moving rats were reported by Segieth et al. (2001). In the rat PAG, 5-HT inhibits, by means of activation of NOS, the pressor response and antinociception produced by local electrical stimulation (Hamalainen and Levick., 1997). In the guinea pig LDTN/PPN, NOSþ neurons are closely interdigitated with 5-HT neurons. The NOSþ neurons receive input from these 5-HT neurons, as well as from NE neurons (which in the guinea pig are not localized into a compact locus ceruleus as they are in the rat) (Leonard et al., 1995). These authors suggest that local NO modulates both THþ and 5-HT neurons in this locality. Another interesting species diVerence is reported by Leger et al. (1998). The raphe nuclei in the rat contain a ‘‘substantial number’’ of NOSþ 5-HT neurons, whereas the raphe in guinea pig and cat do not. IV. Postsynaptic Cascades

All 5-HT receptors, except the 5-HT3R that controls an ionic channel, are linked to G-proteins. The system is one of enormous complexity. Raymond et al., (2001) have published a comprehensive review of this subject. They report the action of the known 5-HT receptors (reactions in nervous tissue marked with *) as follows. A. 5-HT(1A)R Adenylyl cyclase—mostly inhibition, but possibly excitation in the hippocampus* The PI pathway (PLC based)—mostly excitation, but possibly inhibition in the hippocampus* Other phospholipases—activation of phosphatidyl-specific PLC Protein kinases—activation of PKC Proliferation factors—activation of ERK, NFB, and EGF pathways

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Miscellaneous transport processes—stimulation of Naþ/Kþ-ATPase, stimulation of a Naþ/Hþ exchanger B. 5-HT(1B)R Adenylyl cyclase—inhibition* The PI pathway (PLC)—activation Other phospholipases—stimulation of PLD Proliferation factors—stimulation of ERK*, PI-3K, p70 S6 kinase, and Akt kinase C. 5-HT(1D)R Adenylyl cyclase—inhibition* D. 5-HT(1E)R Adenylyl cyclase—inhibition ( low concentrations), stimulation (high concentrations) E. 5-HT(1F)R The PI pathway—activation F. 5-HT(2A)R Adenylyl cyclase—no action in most tissues The PI (PLC) pathway—activation* Other phospholipases—activation of PLA2 and PLD Proliferation factors—ERK stimulation: JaK stimulation Transport processes—Naþ-proton exchanger and Naþ/Kþ-ATPase stimulation G. 5-HT(2B)R Adenylyl cyclase—no activity in most cases The PI (PLC) pathway—activation* Other phospholipases—activation of PLA2

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Proliferation factors—activation of ERK Transport processes—activation of bicarbonate secretion and NaþKþ-ATPase H. 5-HT(2C)R (Confined to Nervous Tissue) Adenylyl cyclase—no activity at physiological levels in most areas* The PI (PLC) pathway—activation* Other phospholipases—activation of PLA2* Proliferation factors—little data Transport processes—activation of several including the Naþ/Ca2þ exchanger* I. 5-HT(4)R (Mostly in Heart and Gut) Adenylyl cyclase—activation (main activity)* J. 5-HT(5)R (Confined to Nervous Tissue) Little data K. 5-HT(6)R (Predominantly in Brain) Adenylyl cyclase—activation* L. 5-HT(7)R (Plentiful in Brain) Adenylyl cyclase—activation* Proliferation factors—activation of ERK*

V. Serotonin and Synaptic Plasticity

Serotonin has manifold eVects on synaptic plasticity. In rodent thalamic cells in culture, 5-HT(1B)Rs induce neurite growth in thalamocortical neurons (Lotto et al., 1999). In rat ventrobasal thalamus, 5-HT promotes growth of neurites as measured by both length and number of branches (Lieske et al., 1999). In rat HC CA1, 5-HT maintains spine density on apical dendrites but is not crucial for steroid-induced synaptic plasticity (raised by estradiol and diminished by

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progesterone) (Alves et al., 2002). In rat dentate gyrus, neonatal 5-HT depletion leads to reduced activity of 5-HT(1A)Rs, which in turn leads to permanent loss of dendritic spines (Yan et al., 1997). In rat dorsomedial PFC layer III pyramidal cells, 5-HT depletion results in shorter dendrites and an increased density of apical and oblique dendrites. There is also an increased number of thin spines on basilar and oblique dendrites and of stubby spines on apical dendrites (Perez-Vega et al., 2000). In rat parietal cortex, synaptic density is increased by 5-HT (but not by DA), whereas the reverse is the case in rat PFC (Sugahara and Shiraishi, 1998). In aplysia sensory neurons, 5-HT (by means of PKA) upregulates the expression of syntaxin in the cell body and promotes its packaging for transport down axons (Hu et al., 2003). In the HC, 5-HT reuptake blockers upregulate the Ca2þ/calmodulin-dependent phosphorylation of synaptotagmin but not of synapsin (Popoli et al., 1997). In rat PFC layer V pyramidal cells, 5-HT(2A)Rs increase glu-evoked EPSPs by means of a late synchronous glu release linked to the high aYnity Ca2þ sensor synaptotagmin. 5-HT also inhibits the early synchronous component of electrically evoked EPSPs (Aghajanian and Marek, 1999a). In rat cerebral cortex, injections of 5-HT(1A)R agonists increase levels of synaptophysin particularly in somata. Injections of the neuronal extension factor S100b did the same but mainly on neuronal processes (Nishi et al., 1996). S100b is an astroglial protein that plays a key role in synaptic production by means of neurite extension, stabilization of microtubules and dendrites, and modulation of GAP–43. 5-HT releases S100b. In rats, 5-HT depletion between postnatal days 10 and 20 leads to loss of MAP-2 and to long-term defects in spatial learning (Mazer et al., 1997). In rat dentate gyrus, 5-HT increases the number of growing neurons and varies the expression of highly polysialylated neural cell adhesion molecule (PSA-NCAM). The authors point out that because 5-HT neurons do not express PSA-NCAM, this modulation must occur in the cellular environment. Astrocytes secrete tenascin C that inhibits the sprouting of DA fibers. (Brezun and Daszuta, 1999; 2000). Neurogenesis occurs in the adult HC and is increased by activity in the cAMP–CREB pathway. This pathway modulates the expression of a rate-limiting enzyme involved in the synthesis of PSA-NCAM (Nakagawa et al., 2002). In Xenopus oocytes, the cell adhesion protein NCAM inhibits the transport by lipid rafts of the GIRK channel Kir3 to the cell surface and thus modulates the long-term excitability of these neurons (Delling et al., 2002). Rat growth cones release 5-HT tonically. This is stimulated by Kþ in a Ca2þdependent manner (Mercado et al., 1998). In rat HC, mineralocorticoids and glucocorticoids modulate in a complex manner 5-HT(1A)R (but not 5-HT(1B)R) mRNA levels (see Neumaier et al., 2000 for details). In cultured embryonic rat (day 14) rostral raphe neurons, BDNF (also cAMP) evokes a dramatic increase in cell numbers and in the growth and branching of dendrites. This is associated with a marked increase in the levels of mRNAs for the 5-HT transporter the

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5-HT(1A)Rs and 5-HT(1B)Rs and the BDNF receptor TrkB (Rumajogee et al., 2002). In rat cerebellar granule cells, 5-HT(2)Rs upregulate levels of 5-HT(2A)r mRNA and its binding sites. This requires de novo synthesis of mRNA and protein by means of activator protein-1 (AP-1) and cAMP-responsive element binding protein (CREB). This process also needs calmodulin and Ca2þ/calmodulindependent kinases (Chalecka-Franaszek et al., 1999; Chen et al., 1995). Activation of 5-HT(2A) and (2C)Rs elevate the expression on mRNA for activity-regulated cytoskeleton-associated protein (Arc). Arc is an immediate early gene selective for dendrites (Pei et al., 2000). A. Serotonin and Scaffolding Proteins As in the case of other receptors, serotonin receptors are tied to scaVolding proteins that align them in proximity to the molecules with which they interact. In NG180-15 cells, 5-HT(3A)Rs are anchored in clusters to F-actin (Emerit et al., 2002). In rat cerebral cortex, 5-HT(2A)Rs are located intracellularly in dendrites (but not in spines) of pyramidal cells in association with MAP1A (Cornea-Herbert et al., 2002). In COS-7 cells, MUPP1 (multi-PDZ domain protein 1), which has 13 PDZ domains, acts as a scaVold for 5-HT(2C)Rs (Be´ camel et al., 2001). B. Serotonin and Learning Serotonin systems play a key role in various types of learning, including long-term potentiation and conditioning. 1. LTP In rat superior cervical ganglion, long-term potentiation (LTP) is dependent on activity in 5-HT(3)Rs and requires NO for maintenance but not for induction. Induction of LTP requires 5-HT(3)R–evoked production of CO as a retromessenger (Alkadhi et al., 2001). In rat HC CA1, 5-HT(4)Rs and ACh MRs cooperate in maintaining LTP. This process involves modulation of spike amplitudes in the CA1 field (Matsumoto et al., 2001). In contrast, in rat dentate gyrus, 5-HT(4) Rs inhibit basic synaptic transmission and LTP (Kulla and Manahan-Vanghan 2002). In a review of the literature, Myhrer (2003) concludes that the multiple memory systems in the rat brain cannot be related to specific transmitter systems because of the great extent of interactions between these systems. 2. Learning and Conditioning In microdialysis studies of the HC in freely moving rats, Wilkinson et al. (1996) showed that 5-HT release is not simply activated by the stimulus but, more subtly, is modulated by whether the adversive stimulus is discrete or contextual

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and also by the magnitude of the conditioning itself. In rat HC CA1, retrieval of inhibitory avoidance memories requires intact gluRs, PKA, PKC, and MAPK in the HC and entorhinal cortex. This system is further modulated by DA(1)Rs, ADRs, muscarinic AchRs, and 5-HT(1A)Rs (Barros et al., 2001). In rat HC (but not amygdala) memory function, the consolidation pathway is dependent on the signal chain cAMP ! PKA and is regulated by dopamine D1Rs, 5-HT(1A)Rs and ADRs (Bevilaqua et al., 1997). In human PET studies, Yasuno et al. (2003) found that 5-HT(1A)Rs in the HC inhibit explicit memory function (as well as lowering body temperature and growth hormone secretion). In mice, retention of learning for foot shock is decreased by 5-HT and GABA agonists and is increased by ACh and glu agonists (Farr et al., 2000). In the rat, conditioned fear responses are associated with raised level of Fosþ cells in the locus coeruleus and dorsal raphe (but not in the VTA) in both NE, 5-HT, and GABA neurons, but not in DA neurons (Ishida et al., 2002). The median raphe nucleus is also concerned with contextual fear conditioning in the rat (Avanzi and Brandao, 2001). In monkey dorsolateral PFC pyramidal cells, 5-HT(2A)Rs increase the spatial timing of ‘‘memory field’’ neurons by increasing activity of preferred target locations and/or reducing activity in nonpreferential locations (Williams et al., 2002). In general, in cognition tasks, 5-HT systems modulate psychological bias (motivation and motor processes), whereas cholinergic systems modulate accuracy. Attention, stimulus processing, and arousal can be modulated by cholinergic and 5-HT systems independently. ACh–5-HT interactions are also important for learning (see review by Steckler and Sahgal, 1995). 3. Backpropagated Potentials (BPPs) When a neuron fires an action potential down its axon, it also sends a backpropagated potential into its dendritic tree. This plays an important role in neuronal Hebbian mechanisms. In rat HC CA1 slices, 5-HT lowers peak potentials of BBPs in apical dendrites by lowering i.c. Ca2þ levels (Sandler and Ross, 1999). In mouse PFC pyramidal cells, 5-HT(2A)Rs and (2C)Rs inhibit BPPs and thus could interfere with Hebbian mechanisms. C. Stress Serotonin also has manifest functions in the stress system. Tailpinch and restraint stress in rat leads to increased levels of 5-HT and 5HIAA in the hippocampus (Vahabzadeh and Fillenz, 1994). In rats, stress (using the communication box paradigm)-induced 5-HT release þþ in the basolateral amygdala and þ in dorsal raphe (as well as DA release þ in basolateral amygdala but not in dorsal raphe) (Funada and Hara, 2001). In rats, overcrowding stress at an early age leads to the loss in both numbers and aYnity of 5-HT(1A)Rs in the HC.

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This is associated with increased anxiety levels. The level of 5-HT in the HC is, however, not aVected (Daniels et al., 2000). Rats reared in isolation from postnatal days 22 to 64 show a loss of 5-HT terminals and of dendrites in HC CA but not in the molecular layer of the dentate gyrus (Whitaker-Azmitia et al., 2000). Wellman (2001) confirmed that stress produces atrophy of apical dendrites in HC CA3 pyramidal cells. Social stress in rats reduces dendritic arborizations in CA3 pyramidal cells by means of the action of 5-HT, glucocorticoids, and other neurotransmitters (McKittrick et al., 2000). Inescapable shock leads to greater increases in activation of raphe neurons and 5-HT eZux in N.Ac. than does escapable shock (Bland et al., 2003). Repeated social stress in rats leads to the atrophy of apical dendrites in the HC CA3 pyramidal cells, which is mediated by glutamate, steroids, and 5-HT. This results in cognitive defects in spatial learning and an increased number of mitochondria (Magarinos et al., 1997). Chronic stress downregulates 5-HT(1A)Rs, and lack of these receptors impairs HC-dependent learning (Sarnyai et al., 2000). In rat HC CA2/3, acute restraint stress increases 5-HT(7)R mRNA levels (Yau et al., 2001), whereas in rat raphe pontis nucleus (but not in other pontine nuclei), acute immobilization stress lowers 5-HT mRNA levels (Vollmayr, 2000). Chronic stress does not aVect 5-HT(7)R levels but lowers mineralocorticoid mRNA levels in the dentate system (Yau et al., 2001). In contrast, in freely moving rats, forced swim stress raises HC 5-HT levels by 1500%. This eVect is dependent on corticotropin-releasing hormone receptor (CRHR) activity (Linthorst et al., 2002). Psychological stress (communication box paradigm) raises constitutive NO production, which results in oxidative damage to membrane lipids. Activation of 5-HT(1A)Rs protects against this type of oxidative stress (Matsumoto et al., 1999). In rats, swim stress activates CRF inputs to dorsal raphe GABA neurons, which in turn inhibit 5-HT neurons and reduce 5-HT release in brain and raise c-fos levels mainly in the GABA neurons (Roche et al., 2003). In the rat, the amygdala facilitates dorsal raphe 5-HT neurons that project to the paraventricular nucleus of the hypothalamus there to modulate the PHA axis. This pathway mediates hypothalamic control of the HPA axis (Weidenfeld et al., 2002). D. Anxiety The HC modulates avoidance components of phobic anxiety, whereas other regions (such as the dorsal hypothalamus) mediate the escape components. In the HC 5-HT(1A)Rs have an anxiogenic action, whereas both nicotinic and muscarinic cholinergic Rs are anxiolytic (File et al., 2000). In studies of anxiety-producing social interaction in rats, 5-HT(1A)Rs in the median raphe were shown to be anxiolytic and in the dorsal HC to be anxiogenic (Andrews et al., 1994). Mutant

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mice lacking 5-HT(1A)Rs show increased anxiety plus tonic disinhibition of central 5-HT transmission, especially in the frontal cortex (Parsons et al., 2001). These mice also have downregulated GABA(A)1 and 2 subunits in the amygdala (Olivier et al., 2001). Activation of the ascending raphe 5-HT pathway to the amygdala and frontal cortex promotes learned defensive behaviors, whereas activation of the pathway from the dorsal raphe to the dorsal PAG inhibits fright and flight reactions (Zangrossi et al., 2001). In rat locus ceruleus, the anxiolytic 5-HT(1A)R agonist azapirone stimulates excitatory 5-HT(1A)Rs on NE neurons (Hamamura et al., 1997).

VI. Serotonin and the Development of Neural Systems

Serotonin is not merely an important neuromodulator in the adult brain but also plays complex roles at various stages of brain development. The details of its system, such as the number and intracortical distribution of its receptors, can show striking changes over time (Kojic et al., 2001). For example, in the superficial (visual) layer of the superior colliculus, 5-HT varicosities form symmetrical synapses with dendrites that increase in number from birth to peak at 1 week postnatal. Their number then falls for a week and then gradually rises again. In contrast, in the deeper (motor) layers of the superior colliculus and in the ventrolateral thalamus, 5-HT varicosities show only a steady rise (Dori et al., 1998). The number of 5-HT(1A)Rs on rat motoneurons reach a peak on postnatal day 7 and then decline to postnatal day 28 (Talley et al., 1997). In rat dentate gyrus, neonatal 5-HT depletion or 5-HT(1A)R blockade leads to a significant and permanent loss of spines because of retardation of synaptogenesis. There is also a compensatory abnormal increase in the number of complex spines (Faber and Haring, 1999). In various layers of the rat primary sensory cortices, there is a transient expression around postnatal day 8 of highaYnity 5-HT reuptake sites (Bennett-Clarke et al., 1996). Selective depletion of noradrenaline and/or serotonin between 1 and 2 weeks after birth decreased the number of synaptic profiles by 29 to 55% compared with that of control animals (Matsukawa et al., 2003). Genetically MAO(A)-deficient mice lack the normal barrel-like clustering of layer IV neurons in the somatosensory cortex. This is due to excessive levels of 5-HT (Cases et al., 1996; Laurent et al., 2002). In the developing barrel-like field of neonatal mice, 5-HT(1B)Rs mediate the inhibition of thalamocortical EPSCs that favor the propagation of high-frequency thalamocortical activity (Laurent et al., 2002). In gerbil superior olive, 5-HT(2)Rs promote prolonged bursts of IPSCs in animals until postnatal day 8 but not thereafter. This eVect is mediated by increasing the excitability of inhibitory aVerents. 5-HT also strongly

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depresses EPSCs by means of 5-HT(1) and 5-HT(2)Rs (Fitzgerald and Sanes, 1999). In rat visual cortical slices, endogenous 5-HT inhibits LTP, and this eVect declines with age (Edagawa et al., 2001). In fetal rat brain, 5-HT promotes the diVerentiation of cortical glu neurons without aVecting the overall population or the percentage of other cell types (Lavdas et al., 1997). In rat raphe cultures, 5-HT(1A)Rs upregulate BDNF to activate its receptors (Trk) to promote serotonergic neuronal development (more neurons) (Galter and Unsicker, 2000). BDNF increases Kþ-stimulated 5-HT release in rat striatal slices by means of its receptor Trk (Goggi et al., 2002). In C6 glioma cells, 5-HT(2A)Rs increase BDNF mRNA expression by means of a Ca2þ and PK-dependent pathway (Meller et al., 2002). GAP-43 is a key regulator for normal pathfinding and arborization of 5-HT axons in early brain development (Donovan et al., 2002). In frog optic tectum, 5-HT(1B) antagonists disrupt the topographical optical map, whereas 5-HT(1A) antagonists refine it (Butt et al., 2002). Gaspar et al. (2003) give a comprehensive review of new genetic information on the role of serotonin in brain development. They conclude that diVerent serotonin receptors, acting at diVerent developmental stages, modulate diVerent developmental processes such as neurogenesis, apoptosis, axon branching, and dendritogenesis.

VII. Sleep

Serotoninergic neurons play a key role in sleep regulation. 5-HT inhibits REM sleep by means of hyperpolarization of LDT neurons (Horner et al., 1997). In the rat, axons from the dorsal raphe 5-HT neurons project to the pedunculopontine nucleus pars compacta, where they synapse on dendrites but not somata (Steininger et al., 1997). The raphe projection to the amygdala in rat also promotes both slow-wave sleep and REM (Gao et al., 2002). In the PPN/LDT nuclei, cholinergic neurons excite glu neurons in the medial pontine reticular formation to promote REM sleep. This eVect is inhibited by 5-HTRs, nicotinic AchRs, and DARs. During REM, GABA(A) neurons inhibit 5-HT neurons in the dorsal raphe. Selective activation of 5-HT(1A)Rs in the dorsal raphe promote REM but in the PPT/LTD inhibit REM (Monti and Monti, 2000). In the cat, dorsal raphe, 5-HT neurons fire tonically— strongly during waking, moderately during slow-wave sleep, and not at all during REM. This inhibition during sleep is due to an increase in GABAergic tone from multiple sources both distant (e. g., the lateral preoptic area during slow-wave sleep and the PAG during REM sleep) and local INs (Gervasoni et al., 2000). In guinea pig LDTN, NOSþ cholinergic neurons increase their tonic rate of firing to drive waking and REM sleep. This is associated with a rise in i.c. Ca2þ levels. 5-HT inhibits the latter both indirectly by reducing cell-firing rates and directly by inhibiting spikeevoked transients. The firing of 5-HT cells is lowest during REM sleep (Leonard

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et al., 2000). PGO-on neurons in the mesopontine tegmentum release more ACh during REM than during waking and slow-wave sleep, and their activity is modulated by 5-HT input (Kodama and Honda, 1996). During waking, 5-HT is released from 5-HT axonal nerve endings, but during sleep, it is released from the dendrites and somata of dorsal raphe cells. The NO synthesis rate is highest in the waking cortex (Cespuglio et al., 1998). In cat mesopontine tegmentum (LTDN/PPNpc), there are two types of cholinergic neurons related to sleep: (1) I-s, which exhibit slow tonic discharge during waking and REM, and (2) PGO-on, which exhibit single spikes during waking and PGO-associated bursts during REM. ACh inhibits single spikes on both types but has no eVect on PGO-burst activity. Glu, NE, epinephrine, and histamine increase tonic discharge in both types. 5-HT has little eVect on either (Koyama and Sakai, 2000). Sleep deprivation in rats leads to reduced levels of neurotransmitters, including serotonin and dopamine, as well as increased dendritic spine counts in the medial preoptic area and motor cortex (Ramesh et al., 1999). The authors point out that the available literature does not indicate a role area for serotonin and dopamine in the medial preoptic area, so that the changes they report may represent their participation in some non-sleep activity.

VIII. Miscellaneous

A. Interleukins In rat C6BU-1 glioma cells, IL-1 inhibits 5-HT(2)R–mediated i.c. Ca2þ mobilization by means of activation of a tyrosine kinase and PKC (Kugaya et al., 1995). In rat anterior hypothalamus, IL-1 acts directly on 5-HT neurons to promote the release of 5-HT (transiently and NE and DA) for a longer time period (Shintani et al., 1993). In mice i.v and i.p., IL-6 activates the HPA axis by upregulating 5-HT synthesis and turnover (Wang and Dunn, 1998). B. Estrogens In rat HC and frontal cortex, estrogens inhibit 5-HT(1A)R function (Mize et al., 2001). C. Opioids In rat medial PFC layer V, pyramidal cells -opioids presynaptically inhibit 5-HT release onto apical dendrites (Marek and Aghajanian, 1998).

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D. Melatonin In chick pineal, prostaglandins (E1 > E2 > F2) increase 5-HT acetylation to form melatonin by means of a Ca2þ/calmodulin-dependent cAMP increase (Voisin et al., 1993). E. Cannabinoids In mouse cortex, cannabinoid CB1Rs inhibit 5-HT release (Nakazi et al., 2000). F. Neurokinin In rat dorsal raphe slices, neurokinins promote 5-HT transmission by raising glu input to raphe neurons (Liu et al., 2002). G. Lysergic Acid Diethylamide (LSD) In rat PFC, LSD is a partial agonist at 5-HT(2A)Rs by means of a Ca2þ/calmodulin-dependent pathway that downregulates NMDA-mediated neurotransmission (Arvanov et al., 1999). In human neuroblastoma cell lines, hallucinogens such as LSD act on 5-HT(2A)Rs but not on 5-HT(2C)Rs (Newton et al., 1996). In rat brain, LSD increases by fivefold to eightfold Fos-like immunoreactivity in the medial PFC, anterior cingulate (wholly by means of 5-HT[2A] Rs), and central nucleus of the amygdala (partly by means of 5-HT[2A]Rs) (but not in the dorsal striatum or nucleus accumbens) (Gresch et al., 2002). Activation by LSD of 5-HT(2A)Rs enhances nonsynchronous, late components of glutamatergic excitatory postsynaptic potentials at apical dendrites of layer V cortical pyramidal cells (Aghajanian and Marek, 1999b). The authors propose that an eVect of hallucinogens on glutamatergic transmission in the cerebral cortex may be responsible for the higher level cognitive, perceptual, and aVective distortions produced by these drugs. H. Reactive Oxygen Species (ROS) 5-HT is neuroprotective in mitochondria by inhibiting ROS (Park et al., 2002) or inhibiting their release (Huether et al., 1997). It is also neuroprotective by free radical scavenging against iron and BSO but not against NMDA

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neurotoxicity (Kang et al., 2001). The antioxidant vitamin ascorbate modulates (upregulates) the binding of 5-HT to its 5-HT3 receptor in a pH-dependent manner (Todd and Bauer, 1988). This indicates that redox factors may play a role in modulation of 5-HT activity (Smythies, 1997, 1999). In rat HC slices, 5-HT stimulates mitochondrial oxidative metabolism as evidenced by an elevation in NAD(P)H fluorescence (Kann et al., 2003). I. Apoptosis In Ca2þ-deficient HN2-5 cells, 5-HT(1A)Rs inhibit apoptosis by two mechanisms (1) by reducing the activity of N-type Ca2þ channels and NMDARs and (2) by MAPK-mediated inhibition of a caspase-3–like enzyme (Adayev et al., 1999). 5-HT KO mice have raised e.c. 5-HT levels and diminished apoptosis (Persico et al., 2003). In chick cultured telencephalic neurons, 5-HT(1A)Rs repress apoptosis by induction of NGF synthesis (Ahlemeyer and Krieglstein, 1997). J. ERK In human transfected neuroblastoma cells, 5-HT(1B)Rs activate e.c. signalregulated kinase (ERK) and protein kinase Akt. This eVect is inhibited by regulator of G-protein signaling protein (RGS) (Leone et al., 2001). In the rat, 5-HT(7)Rs couple to ERK1 and ERK2 pathways. This may be related to the antidepressant eVect of drugs that raise 5-HT levels (Errico et al., 2001). K. Hormones In rat paraventricular nucleus of the hypothalamus, 5-HT(2A)Rs stimulate the release of a number of hormones, i.e., ACTH, oxytocin, renin, and prolactin. They also activate CRF and oxytocin-expressing cells (Van de Kar et al., 2001). L. Peptides In rat visual cortex, 5-HT axons synapse on somatostatin and neuropeptide X containing neurons but never on vasoactive peptide neurons (Paspalas and Papadopoulos, 2001). All CRFþ neurons in the rat dorsa raphe are 5-HTergic. Thus, CRF and 5-HT are co-transmitters in these cells (Commons et al., 2003).

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M. Endocytosis In the NIH 3T3 cell line and in cortical pyramidal cells, the 5-HT(2A)R antagonist clozapine causes 5-HT(2A)Rs to endocytose (Willins et al., 1998). 5-HT(1A)ARs in the rat raphe nucleus are endocytosed but not 5-HT(1A) heteroreceptors in the HC (Riad et al., 2001). N. Aging In rat brain synaptosomes, 5-HT strongly inhibits ACh-stimulated AA release in adult but not in aged brain. 5-HT inhibits the inositol phosphate pathway by means of 5-HT(2)Rs equally in aged and adult brain. However, both 5-HT and ACh more actively stimulate inositol phosphate accumulation in aged than in adult brain (Samochocki and Strosznajder, 1995). O. Dementia In frontotemporal dementia, the 5-HT raphe system, but not the NE locus ceruleus system, is involved ( Yang and Schmitt, 2001). P. Social Dominance Behavior In dominant Anolis carolinensis lizards, dominant males show a more rapid activation of 5-HT in the HC, nucleus accumbens, amygdala, and brainstem during social dominance behavior. However, chronic 5-HT level elevation is associated with submissive behavior (Larson and Summers, 2001). Q. Prepulse Inhibition The PPI is dependent on activity at 5-HT(1A)Rs (Prinssen et al., 2002). R. Beauty and Spirituality Psychotomimetic drugs like LSD act on the serotonin 2A receptor as partial agonists. These drugs can induce ‘‘mystical’’ experiences and hallucinations that have a very powerful esthetic eVect on most people. In other words, people who know about these things and have experienced these hallucinations say that they

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are very beautiful. Therefore, it seems likely that the serotonin system has something to do with esthetic appreciation, although it is diYcult to say how this is aVected. Serotonin also has something to do with a person’s religious outlook. Borg et al. (2003) administered personality tests that measured various parameters to 15 normal male volunteers. The density of 5-HT(1A) receptors was then measured by PET scan. The density levels for the 5-HT(1A)R correlated inversely with the score for self-transcendence but with no other parameter in the tests. The ‘‘self-transcendent’’ parameter reflects whether the subject is open to ideas such as ESP and mysticism in contrast to a reductionistic and empirical world view. The authors concluded that the 5-HT(1A)R is related to spirituality. However, the authors point out that it is not possible on the present evidence to make this link to either low or high activity in this system. This is so, because low binding scores at the 5-HT(1A)R may reflect higher activity at the presynaptic sites where 5-HT(1A)Rs inhibit 5-HT release. It is of interest to note here that LSD aVects 5-HT(2A)Rs, whereas the link that Borg et al. (2003 ) describe is with 5-HT(1A)Rs.

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Wellman, C. L. (2001). Dendritic reorganization in pyramidal neurons in medial prefrontal cortex after chronic corticosterone administration. J. Neurobiol. 49, 245–253. Whitaker-Azmitia, P., Zhou, F., Hobin, J., and Borella, A. (2000). Isolation-rearing of rats produces deficits as adults in the serotonergic innervation of hippocampus. Peptides 21, 1755–1759. Williams, G. V., Rao, S. G., and Goldman-Rakic, P. S. (2002). The physiological role of 5-HT2A receptors in working memory. J. Neurosci. 22, 2843–2854. Williams, S., Serafin, M., Muhlethaler, M., and Bernheim, L. (1998). The serotonin inhibition of high-voltage-activated calcium currents is relieved by action potential-like depolarizations in dissociated cholinergic nucleus basalis neurons of the guinea-pig. Eur. J. Neurosci. 10, 3291–3294. Wirtshafter, D., and Cook, D. F. (1998). Serotonin-1B agonists induce compartmentally organized striatal Fos expression in rats. Neuroreport 9, 1217–1221. Wilkinson, L. S., Humby, T., Killcross, S., Robbins, T. W., and Everitt, B. J. (1996). Dissociations in hippocampal 5-hydroxytryptamine release in the rat following Pavlovian aversive conditioning to discrete and contextual stimuli. J. Neurosci. 8, 1479–1487. Willins, D. L., Alsayegh, L., Berry, S. A., Backstrom, J. R., Sanders-Bush, E., Friedman, L., Khan, N., and Roth, B. L. (1998). Serotonergic antagonist eVects on traYcking of serotonin 5-HT2A receptors in vitro and in vivo. Ann. NY Acad. Sci. 861, 121–127. Wischmeyer, E., and Karschin, A. (1996). Receptor stimulation causes slow inhibition of IRK1 inwardly rectifying Kþ channels by direct protein kinase A-mediated phosphorylation. Proc. Natl. Acad. Sci. USA 93, 5819–5823. Xu, T., and Pandey, S. C. (2000). Cellular localization of serotonin(2A) (5HT(2A)) receptors in the rat brain. Brain Res. Bull. 51, 499–505. Yan, W., Wilson, C. C., and Haring, J. H. (1997). 5-HT1a receptors mediate the neurotrophic eVect of serotonin on developing dentate granule cells. Brain Res. Dev. Brain Res. 98, 185–190. Yan, Q. S., and Yan, S. E. (2001). Serotonin–1B receptor-mediated inhibition of [(3)H]GABA release from rat ventral tegmental area slices. J. Neurochem. 79, 914–922. Yang, Y., and Schmitt, H. P. (2001). Frontotemporal dementia: Evidence for impairment of ascending serotoninergic but not noradrenergic innervation. Immunocytochemical and quantitative study using a graph method. Acta Neuropathol. (Berl.) 101, 256–270. Yasuno, F., Suhara, T., Nakayama, T., Ichimiya, T., Okubo, Y., Takano, A., Ando, T., Inoue, M., Maeda, J., and Suzuki, K. (2003). Inhibitory eVect of hippocampal 5-HT1A receptors on human explicit memory. Am. J. Psychiatry 160, 334–340. Yau, J. L., Noble, J., and Seckl, J. R. (2001). Acute restraint stress increases 5-HT7 receptor mRNA expression in the rat hippocampus. Neurosci. Lett. 309, 141–144. Zangrossi, H., Jr., Viana, M. B., Zanoveli, J., Bueno, C., Nogueira, R. L., and GraeV, F. G. (2001). Serotonergic regulation of inhibitory avoidance and one-way escape in the rat elevated T-maze. Neurosci. Biobehav. Rev. 25, 637–645. Zazpe, A., Artaiz, I., and Del Rio, J. (1994). Role of 5-HT3 receptors in basal and K(þ)-evoked dopamine release from rat olfactory tubercle and striatal slices. Br. J. Pharmacol. 113, 968–972. Zhou, F. M., and Hablitz, J. J. (1999). Activation of serotonin receptors modulates synaptic transmission in rat cerebral cortex. J. Neurophysiol. 82, 2989–2999.

Further Readings

Bagdy, E., Solyom, S., and Harsing, L. G., Jr. (1998). Feedback stimulation of somatodendritic release: A 5-HT3 receptor-mediated eVect in the raphe nuclei of the rat. Brain Res. Bull. 45, 203–208.

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Boeijnga, P. H., and Boddeke, H. W. (1996). Activation of 5-HT1B receptors suppresses low but not high frequency synaptic transmission in the rat subicular cortex in vitro. Brain Res. 721, 59–65. Garabette, M. L., Martin, K. F., and Redfern, P. H. (2000). Circadian variation in the activity of the 5-HT(1B) autoreceptor in the region of the suprachiasmatic nucleus, measured by microdialysis in the conscious freely moving rat. Br. J. Pharmacol. 131, 1569–1576. Kenny, P. J., File, S. E., and Neal, M. J. (2000). Evidence for a complex influence of nicotinic acetylcholine receptors on hippocampal serotonin release. J. Neurochem. 75, 2409–2414. Leone, A. M., Errico, M., Lin, S. L., and Cowen, D. S. (2000). Activation of extracellular signalregulated kinase (ERK) and Akt by human serotonin 5-HT(1B) receptors in transfected BE(2)-C neuroblastoma cells is inhibited by RGS4. J. Neurochem. 75, 934–938. Martin-Ruiz, R., Puig, M. V., Celada, P., Shapiro, D. A., Roth, B. L., Mengod, G., and Artigas, F. (2001). Control of serotonergic function in medial prefrontal cortex by serotonin–2A receptors through a glutamate-dependent mechanism. J. Neurosci. 21, 9856–9866. Matsumoto, M., Togashi, H., Mori, K., Ueno, K., Ohashi, S., Kojima, T., and Yoshioka, M. (2001). Evidence for involvement of central 5-HT(4) receptors in cholinergic function associated with cognitive processes: Behavioral, electrophysiological, and neurochemical studies. J. Pharmacol. Exp. Ther. 296, 676–682. Maura, G., Marcoli, M., Pepicelli, O., Rosu, C., Viola, C., and Raiteri, M. (2000). Serotonin inhibition of the NMDA receptor/nitric oxide/cyclic GMP pathway in human neocortex slices: Involvement of 5-HT(2C) and 5-HT(1A) receptors. Br. J. Pharmacol. 130, 1853–1858. Persico, A. M., Baldi, A., Dell’Acqua, M. L., Moessner, R., Murphy, D. L., Lesch, K. P., and Keller, F. (2003). Reduced programmed cell death in brains of serotonin transporter knockout mice. Neuroreport 14, 341–344. Rodriguez, J. J., Garcia, D. R., and Pickel, V. M. (1999). Subcellular distribution of 5hydroxytryptamine2A and N-methyl-D-aspartate receptors within single neurons in rat motor and limbic striatum. J. Comp. Neurol. 413, 219–231. Sanabria-Bohorquez, S. M., Biver, F., Damhaut, P., Wikler, D., Veraart, C., and Goldman, S. (2002). Quantification of 5-HT(1A) receptors in human brain using p-MPPF kinetic modelling and PET. Eur. J. Nucl. Med. Mol. Imaging. 29, 76–81.

SECTION VI CONCLUSION

The mass of data, and the conclusions that can be drawn from them, does not allow us to formulate any clear-cut overall hypothesis of the function of the neuromodulators covered in this volume. We can only say that the whole system is enormously complex, exhibiting a huge range of intricate and finely engineered anatomical and functional details. Most neuromodulators can either promote or inhibit the activity of neurotransmitters (glutamate and GABA) and of other neuromodulators according to circumstances, such as location, state of the system, diVerent receptors, diVerent timing and concentrations, and diVerent modes of neuronal firing. In some, for example, the nicotinic system, excitation predominates. Except in the case of the cholinergic system in the striatum, the neuromodulators adhere to a common pattern with a small centrally located nucleus, or nuclei, each giving rise to a vast axonal tree that spreads very widely in the brain. This constitutes a very eYcient system, whereby a signal ‘‘X has occurred’’ can be broadcast all over the brain without the need to carry it by individually addressed messages. This saves a vast amount of computational resources. The neuromodulators not only carry the signal ‘‘X has occurred’’ but also modulate the activity of their target neurons so as to deal with X as eYciently as possible. X usually relates to important events of the stimulus field such as novelty, potential dangers or rewards, and salience in general. They modulate certain behavioral reactions to X such as searching the stimulus field for further clues about the implications of X, transferring features of X between diVerent computational and memory mechanisms, and, eventually, deciding on and carrying through a particular strategy to deal with X. The fact that each neuromodulator modulates the eVects of other neuromodulators in so many instances may possibly be explained by the following hypothesis. Because the detection of salience (reward and danger potential) and novelty are behaviorally closely intertwined with each other and with the direction of attention, this may require a lot of cross talk between the neuromodulators mediating these eVects. For example, the interaction between novelty and salience. The brain is continually being bombarded with novel stimuli—I am continually seeing motor cars I have never seen before, etc.—yet I need to attend and react only to novel stimuli with real or potential salience. Suppose that the release of neuromodulator X signals ‘‘novelty’’ and the release of neuromodulator Y signals ‘‘salience.’’ Then the required interaction between salience and novelty may be aVected by computations somewhere in the limbic brain processing the activity of neurons that respond to X and those processing the activity of INTERNATIONAL REVIEW OF NEUROBIOLOGY, VOL. 64 DOI: 10.1016/S0074-7742(05)64006-8

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the neurons responding to Y. But it could also be aVected by presynaptic and postsynaptic cross talk (the action of X on Y’s responses and the action of Y on X’s responses) in individual neurons of the type described extensively in this book. Some interesting new evidence on the role of acetylcholine in the brain has recently emerged. There is now considerable experimental data to show (Kleiser et al., 2004; Kova´ cs et al., 1996; Ramachandran and Blakeslee, 1998; Yarrow et al., 2001) that we normally see not what is actually ‘‘out there’’ but what the brain computes to be most probably ‘‘out there.’’ The visual field that we experience in normal consciousness is a compound structure constructed partly from information coming from the retina by way of the visual thalamocortical projection (‘‘down-up’’ or ‘‘exact inference’’ or I program) and partly from the memorybased ongoing computations by the higher visual brain of what is most probably out there on the basis of its previous experience of these sorts of situations and their development (‘‘up-down’’ or ‘‘naı¨ve inference’’ or P program). The I program is eYcient but expensive in computational resource. The P program is relatively ineYcient but cheaper. Yu and Dayan (2002) suggest that the cholinergic projection from the nucleus basalis to the cortex promotes the I program by two mechanisms. It simultaneously stimulates excitatory nicotinic receptors on the thalamocortical pathway to layer IV cortical neurons, and it also stimulates presynaptic inhibitory muscarinic receptors on the corticocortical pathway in layers I and II that mediates the P program (thus I > P). Thus, when nothing much is happening in the stimulus field, the brain may coast along using more of its P program. Then, if a sudden potentially significant new stimulus occurs, this will stimulate cholinergic neurons in the nucleus basalis that release acetylcholine (ACh) in the cortex. This changes the balance to I > P in the manner described so that computational resources can be focused on the new stimulus and its possible importance estimated (Smythies, 2004). The superior colliculus controls saccadic eye movements often directed at potentially important new stimuli and ACh is also involved in this mechanism. Cholinergic nicotinic input from the pedunculopontine nucleus to the intermediate layer of the superior colliculus activates colliculocortical activity to initiate a saccade (Kobayashi et al., 2002). In seeming contrast, ACh input from the parabigeminal nucleus to the superficial layers of the superior colliculus activates nicotinic receptors on local GABAergic neurons and so inhibits colliculothalamic activity (Lee et al., 2001). In the muscarinic realm, Li et al. (2004) report that activation of presynaptic M1 receptors inhibits GABAergic transmission in the intermediate layer of the SC and so would promote tectofugal transmission. Clearly, the situation in the superior colliculus is more complicated than in the cortex, and further research is needed to clarify the situation. Clearly the Yu and Dayan hypothesis at present applies only to the visual cortex and possibly the prefrontal cortex. There is no evidence at present, as far

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as I know, of a P and an I system in audition. However, the existence of phantom limbs and allied P-type phenomena suggests that such as system also exists robustly in the somatosensory system. The function of the cholinergic system in the motor cortex, limbic system, thalamus, and other subcortical structures remains obscure. There are also interesting correlations between the brain’s I and P systems and similar systems developed in television compression technology (see Smythies, 2005 for details).

References

Kleiser, R., Seitz, R. J., and Krekelberg, B. (2004). Neural correlates of saccadic suppression in humans. Curr. Biol. 14, R195–R197. Kobayashi, Y., Inoue, Y., Yamamoto, M., Isa, T., and Aizawa, H. J. (2002). Contribution of pedunculopontine tegmental nucleus neurons to performance of visually guided saccade tasks in monkeys. J. Neurophysiol. 88, 715–731. Kova´ cs, I., Papathomas, T. V., Yang, M., and Fehe´ r, A. (1996). When the brain changes its mind: Intraocular grouping during retinal rivalry. Proc. Natl. Acad. Sci. USA 93, 508–511. Lee, P. H., Schmidt, M., and Hall, W. C. (2001). Excitatory and inhibitory circuitry in the superficial gray layer of the superior colliculus. J. Neurosci. 21, 8145–8153. Li, F., Endo, T., and Isa, T. (2004). Presynaptic muscarinic acetylcholine receptors suppress GABAergic synaptic transmission in the intermediate grey layer of mouse superior colliculus. Eur. J. Neurosci. 20, 2079–2088. Ramachandran, V. S., and Blakeslee, S. (1998). ‘‘Phantoms in the Brain.’’ Morrow, New York. Smythies, J. (2004). How the brain decides what we see. J. Roy. Soc. Med. 98, 18–20. Smythies, J. (2005). How the brain decides what we see. J. Roy. Soc. Med. 98, 18–20. Yarrow, K., Haggard, P., Heal, R., Brown, P., and Rothwell, J. C. (2001). Illusory perceptions of space and time preserve cross-saccadic perceptual continuity. Nature 414, 302–304. Yu, A. J., and Dayan, P. (2002). Acetylcholine in cortical inference. Neural Networks 15, 719–730.

POSSIBLY OBSCURE ACRONYMS FOR SOME READERS

BDNF BLA BPP DAT E ILN IN LC LDTN MR MS/DB NBM NOS NPY NR OP PSC PB PPN PVN ROS SCN SON TNF VO(A)CC VTA

Brain-derived neurotrophic factor Basolateral amygdala Back-propagated potential Dopamine transporter Epinephrine Intralaminar nuclei Interneuron Locus coereleus Lateral dorsal tegmental nucleus Muscarinic receptor Medial septum/diagonal band Nucleus basalis of Meynart Nitric oxide synthase Neuropeptide Y Nicotinic receptor Organo-phosphate Post-synaptic current Pyridostigmine bromide Pedunculopontine nucleus Paraventricular nucleus Reactive oxygen species Suprachiasmatic nucleus Supraoptic nucleus Tumor necrosis factor Voltage activated (gated) calcium channels Ventral tegmental area

273

INDEX

A AA. See Arachidonic acid Acetylcholine (ACh) dopamine system and, 137–138, 146, 152 in muscarinic system, 17, 18, 19–24, 30–31, 40, 44, 46, 47, 48, 50, 51, 52, 55 nicotinic system and, 55, 57, 61–66, 74, 79 norepinephrine system and, 174–175, 179, 180, 184 Acetylcholinesterase (AChE) in muscarinic system, 42–43, 45 norepinephrine system and, 196–197 ACh. See Acetylcholine AChE. See Acetylcholinesterase AD. See Alzheimer’s disease Adenosine in muscarinic system, 31 Adenylyl cyclase in muscarinic system, 25, 27–28, 32, 38 in serotonin system, 236, 237, 238 Adrenaline system PVN and, 213 SON and, 213 Afterhyperpolarizations (AHP) in serotonin system, 224, 225 Aging serotonin system and, 248 AHP. See Afterhyperpolarizations Alzheimer’s disease (AD) muscarinic system and, 34, 53–54 nicotinic system and, 79–80 norepinephrine system and, 186, 196 AMPA dopamine system and, 133, 150 nicotinic system and, 56, 61, 73–74, 75, 78 AMPAR activation in muscarinic system, 16, 17, 19–20, 21, 31 in norepinephrine system, 182 AMPARs in serotonin system, 228–229

Anatomy of cholinergic system, 2–4 of dopamine system, 124–129 of norepinephrine system, 174–175 of serotonin system, 218–220 Andrenergic receptors (AR) norepinephrine system and, 174–196 Anticholinesterases nicotinic system and, 77 Apoptosis dopamine system and, 148 muscarinic system and, 28, 33, 34, 40–42 nicotinic system and, 77 in serotonin system, 247 AR. See Andrenergic receptors Arachidonic acid (AA) dopamine system and, 144 in muscarinic system, 10, 14, 20, 26, 30, 31, 35 nicotinic system and, 68, 69, 78 Arrestins muscarinic system and, 39 B Backpropagated potentials (BPP) in dopamine system, 146 serotonin system and, 241 BDNF. See Brain-derived neurotrophic factor Behavior dopamine system and, 150–153 muscarinic system and, 51–52 norepinephrine system and, 194–196 serotonin system and, 248 Biphasic effects in norepinephrine system, 177, 181 BPP. See Backpropagated potentials Brain-derived neurotrophic factor (BDNF) dopamine system and, 147 muscarinic system and, 35–36 nicotinic system and, 78–79 275

276

INDEX

Brain-derived neurotrophic factor (BDNF) (cont.) norepinephrine system and, 191–192 serotonin system and, 218, 239, 240, 244 C CA neurons in muscarinic system, 4, 8, 9, 13, 14, 15–17, 34, 40 in serotonin system, 222, 223, 224, 226 Calcineurin norepinephrine system and, 187 Calcium channels in dopamine system, 130–131, 143 in norepinephrine system, 179–180 in serotonin system, 223–227 Calcium currents in muscarinic system, 12–15, 18–19, 27, 35, 37 Calmodulin-kinases (CaMK) dopamine system and, 133, 134, 141, 150 nicotinic system and, 61, 80 CAM. See Cell adhesion molecules CaMK. See Calmodulin-kinases cAMP pathways dopamine system and, 140, 141, 144, 147, 150 norepinephrine system and, 182–183, 184, 185–186, 188–189, 191, 192 serotonin system and, 224, 231, 239 cAMP response-element binding protein (CREB) signaling dopamine system and, 140–141, 142 muscarinic system and, 26 nicotinic system and, 61, 68, 70, 79–80 norepinephrine system and, 185, 186, 191 serotonin system and, 239, 240 Cannabinoids serotonin system and, 246 Carbachol muscarinic system and, 12–13, 31, 34, 35, 52 nicotinic system and, 59, 74 Caveolae muscarinic system and, 37, 39 Cell adhesion molecules (CAM) muscarinic system and, 45 cGMP pathways serotonin system and, 229, 236 Cholecystokinin (CCK) muscarinic system and, 6, 24

Cholinergic neurons in muscarinic system, 5, 6, 7, 18, 22, 52–53 Cholinergic system, 1–2 GABAergic neurons and, 3 hippocampus and, 3, 4 macroanatomy of, 3–4 NBM neurons and, 3, 4 serotonin interaction with, 233–234 striatum and, 2–3, 6 Circadian rhythms muscarinic system and, 54 Consciousness/spirituality serotonin system and, 248–249, 269–270 Cyclooxygenase (COX) norepinephrine system and, 187–188, 192 Cortex muscarinic system and, 17–18, 24, 32, 42 nicotinic system and, 56–57 COX. See Cyclooxygenase CREB. See cAMP response-element binding protein Cyclic nucleotides muscarinic system and, 13, 14, 25, 27–28, 32, 45, 46, 53, 54 Cyclooxygenase (COX) dopamine system and, 153 in muscarinic system, 30–31, 33 in nicotinic system, 69 Cytokines muscarinic system and, 26, 42 nicotinic system and, 78–79 norepinephrine system and, 192 Cytoskeleton dopamine system and, 143 muscarinic system and, 29–30 D DA. See Dopamine DA receptors (DR) muscarinic system and, 22–24, 49 DA release in dopamine system, 124–125, 128, 132–139, 144, 146, 150, 151, 152, 155 DARP. See Dopamine-releasing protein DARPP. See Dopamine/cAMP-regulated phosphoprotein DAT. See Dopamine transporters Dementia serotonin system and, 248

INDEX

Dopamine (DA) muscarinic system and, 16–17, 22–24 nicotinic system and, 64–65, 74 serotonin system and, 219, 230, 231, 234, 241 Dopamine neurons serotonin system and, 223, 235 Dopamine receptors (DR) activation of, 128 distribution of, 124–127 Dopamine system AA and, 144 ACh in, 137–138, 146, 152 AMPA in, 133, 150 anatomy of, 124–129 apoptosis and, 148 backpropagation in, 146 BDNF and, 147 behavior and, 150–153 calcium channels in, 130–131, 143 CaMK and, 133, 134, 141, 150 cAMP pathway and, 140, 141, 144, 147, 150 COX and, 153 CREB and, 140–141, 142 cytoskeleton and, 143 DA release in, 124–125, 128, 132–139, 144, 146, 150, 151, 152, 155 DARP and, 145 DARPP and, 140, 141, 142, 147 DATs in, 145–146, 155 DR activation in, 128 DR distribution in, 124–127 electrophysiology of, 129–132 endocytosis in, 144–145 EPSCs and, 132, 133 ERKs and, 140–141 exocytosis in, 128–129 GABA and, 130, 133, 136–137 glutamate and, 133–136 G-proteins in, 130, 131, 137, 139–140, 149 HC and, 149 learning and, 148–150 LTD and, 148–150 LTP and, 148–150 MAPKs and, 140–141, 142, 147 NE and, 152, 153 neuronal development and, 148 NGF and, 146–147 NMDARs in, 133, 134, 135, 140, 145, 149, 151, 155

277

NO and, 138–139, 150 NT interactions in, 132–139 Parkinson’s disease and, 153 PFC and, 124–125, 128, 129, 130, 132, 133, 137, 145, 146, 149, 150–152 PKs and, 131, 132, 140, 141, 142, 143, 147, 150 plasticity and, 147 potassium channels in, 129–130 prepulse inhibition in, 153 ROS and, 143–144, 153, 154 serotonin interaction with, 234–235 signaling cascades in, 139–143 sodium channels in, 131–132 transmission in, 145–146 tropins and, 146–147 VOCCs and, 130–131 VTA and, 124–125, 127, 129, 132, 135–136, 137, 149, 150 Dopamine transporters (DAT) in dopamine system, 145–146, 155 serotonin system and, 235 Dopamine/cAMP-regulated phosphoprotein (DARPP) dopamine system and, 140, 141, 142, 147 Dopamine-releasing protein (DARP) dopamine system and, 145 Dopaminergic neurons muscarinic system and, 4, 9, 24 E EGF. See Endothelial growth factor Electrical activity in norepinephrine system, 178–180 in serotonin system, 223–227 Electrophysiology of dopamine system, 129–132 Endocytosis in dopamine system, 144–145 in muscarinic system, 22, 32, 37–39 in nicotinic system, 70–71 in norepinephrine system, 190 in serotonin system, 248 Endothelial growth factor (EGF) receptors muscarinic system and, 36 EPSC. See Excitatory postsynaptic currents EPSP. See Excitatory postsynaptic potentials ERK. See Extracellular signal-regulated kinase

278

INDEX

Focal adhesion kinases (FAK) muscarinic system and, 26–27, 33, 36

in muscarinic system, 29 Gene induction in muscarinic system, 44–45 GIRK. See G-protein inwardly rectifying potassium channels Glutamate dopamine system and, 132–136 muscarinic system and, 19–20, 32, 43 nicotinic system and, 61–62 norepinephrine system and, 176–177, 180–182 Glutamate transmission serotonin system and, 228–230, 230–231 G-protein inwardly rectifying potassium channels (GIRK) muscarinic system and, 10, 11, 33, 38 serotonin system and, 224, 225, 239 G-proteins in dopamine system, 130, 131, 137, 139–140, 149 in muscarinic system, 10, 11, 12, 14, 19, 23, 25, 27–28, 29, 30–31, 32, 38, 46, 55 in nicotinic system, 67 in norepinephrine system, 179, 184, 187 in serotonin system, 224, 225, 231, 236

G

H

GABA dopamine system and, 130, 133, 136–137 muscarinic system and, 21–22, 43–44, 52–53 nicotinic system and, 56, 57, 62–63, 64 norepinephrine system and, 175, 182, 183, 184, 185, 186, 194 serotonin system and, 219, 221, 228–229, 231–233, 233–234, 243 GABA transporter 1 (GAT1) muscarinic system and, 22 GABAergic neurons cholinergic system and, 3 muscarinic system and, 4, 5, 6, 9, 13, 16, 17, 18, 19, 21, 22, 52 nicotinic system and, 56, 57, 59, 63, 76 in serotonin system, 219, 222, 224, 231, 232, 241, 242 Gap junctions in norepinephrine system, 197 GAT1. See GABA transporter 1 Gene expression pathways

HC. See Hippocampus Hebbian synapses nicotinic system and, 73–75 Hippocampus (HC) cholinergic system and, 3, 4 dopamine system and, 149 muscarinic system and, 4, 5, 9, 11–13, 15–17, 32, 35–36, 42, 43, 48 nicotinic system and, 56–57, 62, 64, 68, 76 norepinephrine system and, 174–175, 177, 178, 182, 188 serotonin system and, 220, 221, 223, 227, 236, 238–239, 240–241, 242 Histamine norepinephrine system and, 198 Hormones muscarinic system and, 45–46 serotonin system and, 247 5-HT (serotonin) norepinephrine system and, 174–175, 176, 177, 182, 184, 185, 186, 192, 196

Estrogens serotonin system and, 245 Excitatory effects in norepinephrine system, 176 Excitatory postsynaptic currents (EPSC) in dopamine system, 132, 133 in muscarinic system, 10, 19, 21, 31 in nicotinic system, 59, 61, 62, 66 Excitatory postsynaptic potentials (EPSP) in nicotinic system, 72, 76 in serotonin system, 225, 226, 228, 229, 239, 243, 244 Exocytosis in dopamine system, 128–129 in norepinephrine system, 186 Extracellular signal-regulated kinase (ERK) dopamine system and, 140–141 muscarinic system and, 8–9, 25, 26, 41, 49 nicotinic system and, 70, 72, 79–80 norepinephrine system and, 186–187 serotonin system and, 236, 237, 238, 247 F

INDEX

5-HT neurons in serotonin system, 218–220, 228, 230, 237 5-HT release in serotonin system, 221, 227, 245 5-HT1 receptors in serotonin system, 220–221, 223 5-HT2 receptors in serotonin system, 221–222, 223 I Inhibitory effects in norepinephrine system, 176–177 Inhibitory postsynaptic potentials (IPSP) in serotonin system, 226, 228, 230, 231, 243 Inositol 1,4,5-triphosphate (IP3) signaling in muscarinic system, 14, 15, 28 Integrins muscarinic system and, 10, 11, 27, 45 Interleukins serotonin system and, 245 Ionic conductance in nicotinic system, 60–61 IP3. See Inositol 1,4,5-triphosphate IPSP. See Inhibitory postsynaptic potentials K Knockout mice muscarinic system and, 10, 12, 22 L LC. See Locus ceruleus Learning dopamine system and, 148–150 muscarinic system and, 48–51 nicotinic system and, 72–77 norepinephrine system and, 192–196 serotonin system and, 240–241 Locus ceruleus (LC) norepinephrine system and, 174–177, 181, 182, 183, 184 serotonin system and, 229, 243 Long-term depression (LTD) dopamine system and, 148–150 norepinephrine system and, 192–193 Long-term potentiation (LTP) dopamine system and, 148–150 muscarinic system and, 48, 49, 50

279

nicotinic system and, 72–73 norepinephrine system and, 192–196 serotonin system and, 240, 244 Low-voltage fast activity (LVFA) muscarinic system and, 17–18 LSD. See Lysergic acid diethylamide LTD. See Long-term depression LTP. See Long-term potentiation LVFA. See Low-voltage fast activity Lysergic acid diethylamide (LSD) serotonin system and, 246 M MAPK. See Mitogen-activated protein kinases Melatonin serotonin system and, 246 Mitogen-activated protein kinases (MAPK) in dopamine system, 140–141, 142, 147 in muscarinic system, 9, 24–26, 41, 44, 49 in nicotinic system, 61, 67, 70 in norepinephrine system, 186–187 in serotonin system, 225, 241, 247 mRNAs in nicotinic system, 57–58 in serotonin system, 220, 222, 239–240, 242, 244 Muscarinic 1 receptors (M1R) in muscarinic system, 4–5, 13, 18, 22, 24–26, 51 Muscarinic 2 receptors (M2R) in muscarinic system, 3, 5–7, 17, 22, 31, 35, 38, 42 Muscarinic receptors (MR) activation of, 9, 10, 12, 13, 14, 15, 17, 20, 21, 30, 36 Muscarinic system AA in, 10, 14, 20, 26, 30, 31, 35 ACh in, 17, 18, 19–24, 30–31, 40, 44, 46, 47, 48, 50, 51, 52, 55 AChE in, 42–43, 45 AD and, 34, 53–54 adenosine in, 31 adenylyl cyclase in, 25, 27–28, 32, 38 AMPAR activation in, 16, 17, 19–20, 21, 31 apoptosis and, 28, 33, 34, 40–42 arrestins and, 39 BDNF and, 35–36 behavior and, 51–52 CA neurons and, 4, 8, 9, 13, 14, 15–17, 34, 40

280

INDEX

Muscarinic system (cont.) calcium currents in, 12–15, 18–19, 27, 35, 37 CAMs and, 45 carbachol and, 12–13, 16, 31, 34, 35, 52 caveolae in, 37, 39 CCKs and, 6, 21, 24 cholinergic neurons and, 5, 6, 7, 18, 22, 52–53 circadian rhythms and, 54 cortex and, 17–18, 24, 32, 42 COX in, 30–31, 33 CREB signaling and, 25, 26 cyclic nucleotides in, 13, 14, 25, 27–28, 32, 45, 46, 53, 54 cytokines and, 26, 42 cytoskeleton and, 29–30 DA and, 16–17, 22–24 DA receptors and, 22–24, 49 dopaminergic neurons and, 4, 9, 24 EGF receptors in, 36 endocytosis in, 22, 32, 37–39 EPSCs in, 10, 19, 21, 31 ERKs in, 8–9, 25, 26, 41, 49 FAKs and, 26–27, 33, 36 GABA and, 21–22, 43–44, 52–53 GABAergic neurons and, 4, 5, 6, 9, 13, 16, 17, 18, 19, 21, 22, 52 GAT1 and, 22 gene expression in, 29 gene induction in, 44–45 GIRKs and, 10, 11, 33, 38 glutamate and, 19–20, 32, 43 G-proteins in, 10, 11, 12, 14, 19, 23, 25, 27–28, 29, 30–31, 32, 38, 46, 55 HC and, 4, 5, 9, 11–13, 15–17, 32, 35–36, 42, 43, 48 hormones and, 45–46 integrins and, 10, 11, 27, 45 IP3 signaling in, 14, 15, 28 knockout mice in, 10, 12, 22 learning and, 48–51 LTP and, 48, 49, 50 LVFA and, 17–18 M1Rs in, 4–5, 13, 18, 22, 24–26, 51 M2Rs in, 3, 5–7, 17, 22, 31, 35, 38, 42 MAPKs in, 9, 24–26, 41, 44, 49 MR activation in, 9, 10, 12, 13, 14, 15, 17, 20, 21, 30, 36 NBM in, 4–5, 35, 36, 48, 49 neuroprotective effects in, 41–42 neurotoxic effects in, 40–41, 42–44

NGF and, 35–36, 53 NMDAR activation in, 16, 17, 19, 20, 21, 23, 30, 31, 33, 34, 37, 40–41, 43, 48–50 NOS in, 7, 30, 33, 34, 35, 39, 54 oscillatory activity in, 15–18 oxytocin in, 46–47 PG synthesis in, 31 phospholipases and, 27–28, 30–31, 39–40, 41 PI signaling in, 33, 35, 38, 39, 41, 49 PKs in, 8–9, 11, 13, 21, 22, 23, 25, 26, 30, 37, 44, 45, 54 plasticity in, 44 PLC signaling in, 24, 27–28, 29, 37, 38, 39, 46, 55 polarization states in, 9–10, 12, 13, 14, 15–17, 18, 26–27 post-synaptic cascades and, 24–28 potassium currents in, 8–11, 15–16, 18, 22, 27, 33 PTX inhibitor and, 12, 25, 27–28, 33 PYKs and, 26–27 pyramidal neurons and, 6, 9, 12, 14, 15–16, 18, 19, 48 redox factors and, 33–34 ROS in, 30–31, 32–34, 40–41 signaling pathways in, 10–11 sodium currents in, 11–12, 15, 18 Src signaling in, 20, 21, 25, 27 stress and, 51 striatum and, 6, 7, 12, 18, 20, 32, 40 t-cells and, 48 transferrins and, 39 tropins and, 35–37 tyrosine phosphorylation in, 26–27 vasodilation and, 7 vasopressin in, 46–47 VTA and, 23, 24, 52 N N–methyl-D-aspartate (NMDA) nicotinic system and, 59, 73, 74, 75 serotonin system and, 230, 231, 236, 246 NBM. See Nucleus basalis of Meynert NE. See Norepinephrine Nerve growth factor (NGF) dopamine system and, 146–147 muscarinic system and, 35–36, 53 nicotinic system and, 78–79 Neurokinin

INDEX

serotonin system and, 246 Neuronal development dopamine system and, 148 in norepinephrine system, 193 serotonin system and, 243–244 Neuronal oscillation in serotonin system, 227 Neuroprotection of muscarinic system, 41–42 of norepinephrine system, 189–190 Neurotoxicity of muscarinic system, 40–41, 42–44 in norepinephrine system, 189–190 Neurotransmitter (NT) interactions in dopamine system, 132–139 in norepinephrine system, 180–185 NGF. See Nerve growth factor Nicotine nicotinic system and, 65–66, 67, 68, 70, 76–77 Nicotinic receptor (NR) activation nicotinic system and, 68, 69 Nicotinic receptors (NR) nicotinic system and, 55, 56, 57, 62, 63, 79 Nicotinic system AA in, 68, 69, 78 ACh in, 55, 57, 61–66, 74, 79 AD and, 79–80 AMPA synapses and, 56, 61, 73–74, 75, 78 anticholinesterases and, 77 apoptosis and, 77 BDNF and, 78–79 CaMKs in, 61, 80 carbachol and, 59, 74 cortex and, 56–57 COX in, 69 CREBs in, 61, 68, 70, 79–80 cytokines and, 78–79 DA and, 64–65, 74 endocytosis and, 70–71 EPSCs in, 59, 61, 62, 66 EPSPs in, 72, 76 ERKs in, 70, 72, 79–80 GABA in, 56, 57, 62–63, 64 GABAergic neurons and, 56, 57, 59, 63, 76 glutamate in, 61–62 G-proteins in, 67 HC and, 56–57, 62, 64, 68, 76 Hebbian synapses and, 73–75 ionic conductances in, 60–61

281

learning and, 72–77 LTP in, 72–73 MAPKs in, 61, 67, 70 mRNA distribution in, 57–58 NE and, 63–64 NGF and, 78–79 nicotine and, 65–66, 67, 68, 70, 76–77 NMDA and, 59, 73, 74, 75 NMDAR activation and, 56, 61, 67, 68, 73–75, 76 NO synthesis and, 64, 68, 69–70 NR activation and, 68, 69 NRs and, 55, 56, 57, 62, 63, 79 oxytocin and, 77 PGs and, 69 PKs in, 64, 65, 69, 74, 77, 79–80 plasticity in, 66–67 postsynaptic signaling in, 67–68 ROS in, 69–70 scaffolding proteins and, 71 serotonin and, 65–66 transmitter interactions in, 61–66 tropins and, 78–79 vasopressin and, 77 VOCCs in, 60–61, 68, 70 VTAs in, 64, 68 Nitric oxide (NO) synthesis in dopamine system, 138–139, 150 in muscarinic system, 7, 30, 33, 34, 35, 39, 54 in nicotinic system, 64, 68, 69–70 in norepinephrine system, 186, 188–189, 192, 196, 198 in serotonin system, 235, 240, 242, 245 NMDA, See N–methyl–D–aspartate NMDAR activation in dopamine system, 133, 134, 135, 140, 145, 149, 151, 155 in muscarinic system, 16, 17, 19, 20, 21, 23, 30, 31, 33, 34, 37, 40–41, 43, 48–50 in nicotinic system, 56, 61, 67, 68, 73–75, 76 in norepinephrine system, 176, 181–182, 193 in serotonin system, 231–222, 228, 229, 230, 231 Norepinephrine (NE) dopamine system and, 152, 153 nicotinic system and, 63–64 norepinephrine system and, 175–176, 182, 184, 188, 191, 196 Norepinephrine system ACh in, 174–175, 179, 180, 184

282

INDEX

Norepinephrine system (cont.) AChE in, 196–197 AD and, 186, 196 AMPARs and, 182 anatomy of, 174–175 ARs and, 174–196 BDNF and, 191–192 behavior and, 194–196 biphasic effects in, 177, 181 calcineurin and, 187 calcium channels in, 179–180 cAMP pathways and, 182–183, 184, 185–186, 188–189, 191, 192 COX and, 187–188, 192 CREB and, 185, 186, 191 cytokines and, 192 electrical activity in, 178–180 endocytosis in, 190 ERK and, 186–187 excitatory effects in, 176 exocytosis in, 186 GABA and, 175, 182, 183, 184, 185, 186, 194 gap junctions and, 197 glutamate and, 176–177, 180–182 G-proteins and, 179, 184, 187 HC and, 174–175, 177, 178, 182, 188 histamine and, 198 5-HT and, 174–175, 176, 177, 182, 184, 185, 186, 192, 196 inhibitory effects in, 176–177 LC and, 174–177, 181, 182, 183, 184 learning and, 192–196 LTD and, 192–193 LTP and, 192–196 MAPK and, 186–187 NE release and, 175–176, 182, 184, 188, 191, 196 neuronal development in, 193 neuroprotection in, 189–190 neurotoxicity in, 189–190 NMDARs and, 176, 181–182, 193 NO and, 186, 188–189, 192, 196, 198 oxytocin and, 197 PFC and, 174, 192, 194 PGs and, 186, 187–188 PI pathway and, 179–180, 187 PKs and, 180–181, 185, 186, 190, 198 PLs and, 187, 190 postsynaptic cascades and, 185–191 potassium channels in, 178

PVN and, 188, 192, 193–194, 195, 197 ROS and, 184, 189, 196, 198 serotonin interaction with, 219–220, 235 sodium channels in, 196–197 stress and, 193–194 transmitter interactions in, 180–185 transporters in, 197–198 tropins and, 191–192 vasopressin and, 197 VOCCs and, 178, 179, 180, 196–197 Nucleus basalis of Meynert (NBM) cholinergic system and, 3, 4 muscarinic system and, 3, 35, 36, 48, 49 O Opioids serotonin system and, 245 Oscillatory activity in muscarinic system, 15–18 Oxytocin muscarinic system and, 46–47 nicotinic system and, 77 norepinephrine system and, 197 P Paraventricular nuclei (PVN) adrenaline system and, 213 norepinephrine system and, 188, 192, 193–194, 195, 197 Parkinson’s disease dopamine system and, 153 Peptides serotonin system and, 247 Pertussis toxin (PTX) inhibitor muscarinic system and, 6, 12, 25, 27–28, 33 PFC. See Prefrontal cortex PG. See Prostaglandins Phosphatidylinositol (PI) signaling muscarinic system and, 33, 35, 38, 39, 41, 49 norepinephrine system and, 179–180, 187 serotonin system and, 236, 237, 238 Phosphokinases (PK) in dopamine system, 131, 132, 140, 141, 142, 143, 147, 150 in muscarinic system, 8–9, 11, 13, 21, 22, 23, 25, 26, 30, 37, 44, 45, 54 in nicotinic system, 64, 65, 69, 74, 77, 79–80

INDEX

in norepinephrine system, 180–181, 185, 186, 190, 198 in serotonin system, 224, 225, 226, 227, 229, 232, 236, 241, 244, 245 Phospholipase C (PLC) signaling muscarinic system and, 24, 27–28, 29, 37, 38, 39, 46, 55 serotonin system and, 226, 236, 237, 238 Phospholipases (PL) muscarinic system and, 27–28, 30–31, 39–40, 41 norepinephrine system and, 187, 190 PK. See Phosphokinases PL. See Phospholipases Plasticity in dopamine system and, 147 in muscarinic system, 44 in nicotinic system, 66–67 serotonin system and, 239–240 PLC. See Phospholipase C Polarization states in muscarinic system, 9–10, 12, 13, 14, 15–17, 18, 26–27 Postsynaptic cascades muscarinic system and, 24–28 in norepinephrine system, 185–191 serotonin system and, 236–238 Postsynaptic excitation in serotonin system, 232 Postsynaptic facilitation in serotonin system, 229–230 Postsynaptic inhibition in serotonin system, 229–230, 232 Postsynaptic signaling nicotinic system and, 67–68 Potassium channels in dopamine system, 129–130 in norepinephrine system, 178 in serotonin system, 223–225 Potassium currents in muscarinic system, 8–11, 15–16, 18, 22, 27, 33 Prefrontal cortex (PFC) dopamine system and, 124–125, 128, 129, 130, 132, 133, 137, 145, 146, 149, 150–152 norepinephrine system and, 174, 191, 194 serotonin system and, 221, 222, 223, 226, 229, 231, 232, 239, 241 Prepulse inhibition

283

in dopamine system, 153 serotonin system and, 248 Presynaptic excitation in serotonin system, 231 Presynaptic facilitation in serotonin system, 229 Presynaptic inhibition in serotonin system, 228–229, 231 Proline-rich kinases (PYK) muscarinic system and, 26–27 Prostaglandins (PG) muscarinic system and, 31 nicotinic system and, 69 norepinephrine system and, 186, 187–188 PTX. See Pertussis toxin PVN. See Paraventricular nuclei PYK. See Proline-rich kinases Pyramidal neurons muscarinic system and, 6, 9, 12, 14, 15–16, 18, 19, 48 serotonin system and, 221, 226, 228, 233, 242 R Reactive oxygen species (ROS) dopamine system and, 143–144, 153, 154 muscarinic system and, 30–31, 32–34, 40–41 nicotinic system and, 69–70 norepinephrine system and, 184, 189, 196, 198 serotonin system and, 246–247 Redox factors muscarinic system and, 33–34 ROS. See Reactive oxygen species S Scaffolding proteins nicotinic system and, 71 Serotonin. See also 5-HT nicotinic system and, 65–66 Serotonin system 5-HT neurons in, 218–220, 228, 230, 237 5-HT release in, 221, 227, 245 5-HT1 receptors in, 220–221, 223 5-HT2 receptors in, 221–222, 223 adenylyl cyclase pathway in, 236, 237, 238 aging and, 248 AHPs in, 224, 225 AMPARs in, 228–229

284 Serotonin system (cont.) anatomy of, 218–220 apoptosis and, 247 BDNF and, 218, 239, 240, 244 behavior and, 248 BPPs and, 241 CA neurons and, 222, 223, 224, 226 calcium channels in, 223–227 cAMP and, 224, 231, 239 cannabinoids and, 246 cGMP and, 229, 236 cholinergic system and, 233–234 consciousness/spirituality and, 248–249, 269–270 CREB and, 239, 240 DA and, 219, 230, 231, 234, 241 DA neurons and, 223, 235 DATs and, 235 dementia and, 248 dopamine system and, 234–235 electrical actions in, 223–227 endocytosis and, 248 EPSPs in, 225, 226, 228, 229, 239, 243, 244 ERKs and, 236, 237, 238, 247 estrogens and, 245 GABA in, 219, 221, 228–229, 231–233, 233–234, 243 GABAergic neurons in, 219, 222, 224, 231, 232, 241, 242 GIRKs and, 224, 225, 239 glutamate transmission and, 228–230, 230–231 G-proteins and, 224, 225, 231, 236 HC and, 220, 221, 223, 227, 236, 238–239, 240–241, 242 hormones and, 247 interleukins and, 245 IPSPs in, 226, 228, 230, 231, 243 LCs and, 229, 243 learning and, 240–241 LSD and, 246 LTP and, 240, 244 MAPKs and, 225, 241, 247 melatonin and, 246 mRNAs in, 220, 222, 239–240, 242, 244 NE and, 219–220 neurokinin and, 246 neuronal development and, 243–244 neuronal oscillation in, 227 NMDA and, 230, 231, 236, 246

INDEX

NMDARs and, 221–222, 228, 229, 230, 231 NO system and, 235, 236, 240, 242, 245 norepinephrine interaction with, 235 NT interactions with, 228–236 opioids and, 245 peptides and, 247 PFC and, 221, 222, 223, 226, 229, 231, 232, 239, 241 PI pathway in, 236, 237, 238 PKs and, 224, 225, 226, 227, 229, 232, 236, 241, 244, 245 plasticity and, 239–240 PLCs and, 226, 236, 237, 238 postsynaptic cascades and, 236–238 postsynaptic excitation in, 232 postsynaptic facilitation in, 229–230 postsynaptic inhibition in, 229–230, 232 potassium channels in, 223–225 prepulse inhibition and, 248 presynaptic excitation in, 231 presynaptic facilitation in, 229 presynaptic inhibition in, 228–229, 231 pyramidal neurons and, 221, 226, 228, 233, 242 ROS and, 246–247 sleep/REM and, 233, 244–245 stress/anxiety and, 241–243 striatum and, 220–221 synaptic v. volume transmission in, 223 VOCCs and, 229 VTA and, 220, 231, 235 Sleep/REM serotonin system and, 233, 244–245 Sodium channels in dopamine system, 131–132 in norepinephrine system, 196–197 Sodium currents in muscarinic system, 11–12, 15, 18 SON. See Supraoptic nuclei Src signaling in muscarinic system, 20, 21, 25, 27 Stress muscarinic system and, 51 norepinephrine system and, 193–194 serotonin system and, 241–243 Striatum cholinergic system and, 2–3, 6 muscarinic system and, 6, 7, 12, 18, 20, 32, 40

INDEX

serotonin system and, 220–221 Supraoptic nuclei (SON) adrenaline system and, 213 T T-cells muscarinic system and, 48 Transferrins muscarinic system and, 39 Transmission in dopamine system, 145–146 Transmitter interactions in nicotinic system, 61–66 Transporters in norepinephrine system, 197–198 Tropins dopamine system and, 146–147 muscarinic system and, 35–37 nicotinic system and, 78–79 norepinephrine system and, 191–192 Tyrosine phosphorylation in muscarinic system, 26–27

285

V Vasodilation muscarinic system and, 7 Vasopressin muscarinic system and, 46–47 nicotinic system and, 77 norepinephrine system and, 197 Ventral tegmental area (VTA) dopamine system and, 124–125, 127, 129, 132, 135–136, 137, 149, 150 muscarinic system and, 23, 24, 52 nicotinic system and, 64, 68 serotonin system and, 220, 231, 235 VOCC. See Voltage-activated calcium channels Voltage-activated calcium channels (VOCC) in dopamine system, 130–131 in nicotinic system, 60–61, 68, 70 in norepinephrine system, 178, 179, 180, 196–197 in serotonin system, 229 Volume transmission in serotonin system, 223 VTA. See Ventral tegmental area

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The Neuromodulators, (International Review of Neurobiology)

INTERNATIONAL REVIEW NEUROBIOLOGY

The Neurobiology of Painting, Volume 74: International Review of Neurobiology

The Neurobiology of Painting - International Review of Neurobiology Vol 74

International Review of Neurobiology, Volume 6

International Review of Neurobiology Volume 25

International Review of Neurobiology Volume 38

International Review of Neurobiology Volume 18

International Review of Neurobiology Volume 31

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International Review of Neurobiology, Volume 23

International Review of Neurobiology Volume 1

International Review of Neurobiology Volume 13

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International Review of Neurobiology, Volume 21

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DNA Arrays in Neurobiology (International Review of Neurobiology, Vol 60)

International Review of Neurobiology Volume 17

International Review of Neurobiology Volume 4

International Review of Neurobiology, Volume 63

International Review of Neurobiology Volume 2

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