Glutamate (Handbook of Chemical Neuroanatomy)

GLUTAMATE

GLUTAMATE

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H A N D B O O K OF CHEMICAL NEUROANATOMY Series Editors" A. Bj6rklund and T. H6kfelt

Volume 18

GLUTAMATE Editors:

O.R OTTERSEN and J. STORM-MATHISEN Department of Anatomy, Institute of Basic Medical Sciences, University of Oslo, RO. Box 1105, Blindern, N-0317 Oslo, Norway

2000

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List of Contributors L. BRODIN (p. 273) Department of Neuroscience Nobel Institute for Neurophysiology Karolinska Institutet S- 171 77 Stockholm Sweden lennart.brodin @neuro.ki, se

B. HASSEL (p. 1) Division of Environmental Toxicology Norwegian Defense Research Establishment R O. Box 25 N-2027 Kjeller Norway

J. BROMAN (p. 1) Department of Physiological Sciences Lund University S61vegatan 19 S-223 62 Lund Sweden j onas.broman @mphy.lu, se

T. KANEKO (p. 203) Department of Morphological Brain Science Graduate School of Medicine Kyoto University Kyoto 606-8501 Japan kaneko @mbs.kyoto-u, ac.jp

N.C. DANBOLT (p. 231) Department of Physiology Institute of Basic Medical Sciences University of Oslo RO. Box 1103, Blindern N-0317 Oslo Norway n.c.danbolt @basalmed.uio.no

A. MATSUBARA (p. 255) Department of Otorhinolaryngology Hirosaki University School of Medicine 5 Zaifu-cho Hirosaki 036-8562 Japan

S. FUJITA (p. 255) Department of Otorhinolaryngology Hirosaki University School of Medicine 5 Zaifu-cho Hirosaki 036-8562 Japan

N. MIZUNO (p. 63) Tokyo Metropolitan Institute for Neuroscience Musashidai 2-6 Fuchu Tokyo 183-8526 Japan [email protected]

V. GUNDERSEN (p. 45) Department of Anatomy Institute of Basic Medical Sciences University of Oslo RO. Box 1105, Blindern N-0317 Oslo Norway [email protected]

H. MONYER (p. 99) Department of Clinical Neurobiology University Hospital of Neurology Im Neuenheimer Feld 364 D-69120 Heidelberg Germany monyer@ otto.mpimf-heidelberg.mpg.de

O.P. OTTERSEN (pp. 1,255) Department of Anatomy Institute of Basic Medical Sciences University of Oslo EO. Box 1105, Blindern N-0317 Oslo Norway o.p.ottersen @basalmed.uio.no R.S. PETRALIA (p. 145) Laboratory of Neurochemistry 36/5D08, NIDCD/NIH 36 Convent Drive, MSC 4162 Bethesda, MD 20892-4162 USA petralia @pop.nidcd.nih.gov E. RINVIK (p. 1) Department of Anatomy Institute of Basic Medical Sciences University of Oslo P.O. Box 1105, Blindern N-0317 Oslo Norway [email protected] M.E. RUBIO (p. 145) Max-Planck-Institute for Experimental Medicine Department of Molecular Biology of Neuronal Signals Hermann-Rein-Strasse 3 D-37075 G6ttingen Germany mrubio @gwdg.de P.H. SEEBURG (p. 99) Max-Planck-Institute for Medical Research Department of Molecular Neurobiology Jahnstrasse 29 D-69120 Heidelberg Germany seeburg @otto.mpimf-heidelberg.mpg.de

vi

M. SHENG (p. 183) Department of Neurobiology, HHMI Massachusetts General Hospital 50 Blossom Street (Wellman 423) Boston, MA 02114 USA sheng @helix.mgh.harvard.edu R. SHIGEMOTO (p. 63) Laboratory of Cerebral Structure National Institute for Physiological Sciences Myodaiji, Okazaki 444-8585 Japan shi gemot @nips. ac .jp O. SHUPLIAKOV (p. 273) Department of Neuroscience Nobel Institute for Neurophysiology Karolinska Institutet S- 171 77 Stockholm Sweden oleg. shupliakov @neuro.ki, se J. STORM-MATHISEN (p. 45) Department of Anatomy Institute of Basic Medical Sciences University of Oslo P.O. Box 1105, Blindern N-0317 Oslo Norway j on. storm-mathisen @basalmed.uio.no Y. TAKUMI (p. 255) Department of Otorhinolaryngology Hirosaki University School of Medicine 5 Zaifu-cho Hirosaki 036-8562 Japan

S. USAMI (p. 255) Department of Otolaryngology Shinshu University School of Medicine 3-1-1 Asahi Matsumoto 390-8621 Japan usami @md. shinshu-u, ac.jp Y.-X. WANG (p. 145) Laboratory of Neurochemistry 36/5D08, NIDCD/NIH 36 Convent Drive, MSC 4162 Bethesda, MD 20892-4162 USA wang @nidcd.nih.gov R.J. WENTHOLD (p. 145) Laboratory of Neurochemistry 36/5D08, NIDCD/NIH 36 Convent Drive, MSC 4162 Bethesda, MD 20892-4162 USA wenthold @nidcd.nih.gov

W. WISDEN (p. 99) MRC Laboratory of Molecular Biology MRC Centre Hills Road Cambridge CB2 2QH UK and

Department of Clinical Neurobiology University Hospital of Neurology Im Neuenheimer Feld 364 D-69120 Heidelberg Germany wwl @mrc-lmb.cam.ac.uk M. WYSZYNSKI (p. 183) Department of Neurobiology HHMI, Massachusetts General Hospital 50 Blossom Street (Wellman 423) Boston, MA 02114 USA wyszynski @helix.mgh.harvard.edu

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Preface In the years that have elapsed since glutamate was first reviewed in this book series (Ottersen and Storm-Mathisen, Handbook of Chemical Neuroanatomy, Vol. 3, 1984, pp. 141246) the field of glutamate neurochemistry has changed dramatically. In 1984, glutamate immunocytochemistry was still in its early days, and tracing with the metabolically inert glutamate analogue, D-aspartate, was one of the very few approaches that were available for the identification of putative glutamatergic pathways. Major advances were made in the late 1980s and early 1990s. The adaptation of quantitative immunogold procedures permitted "transmitter pools" of glutamate to be distinguished from "metabolic pools", and the cloning of glutamate receptors was soon followed by generation of specific antibodies. With these tools in hand it became possible to identify sites of glutamate neurotransmission with a high degree of confidence and precision. Ample experimental support could thus be provided of the notion that glutamate mediates signaling in a majority of the synapses in the brain. This notion dates back to the work of Curtis and Watkins (1960, J Neurochem 6:117-141) who observed that sensitivity to the excitatory effects of glutamate was a property common to most neurons. In hindsight, it is amusing to note that this seemingly non-selective action was one reason for the initial reluctance to accept glutamate as a neurotransmitter. With the realization that glutamate is likely to act as a transmitter (or cotransmitter) in most excitatory synapses in the brain, the interest has turned from mapping of pathways to analysis of the "chemical neuroanatomy" of individual glutamate synapses. This shift of focus is duly reflected in the present volume. Thus, whereas Chapter 1 provides an overview of major glutamatergic fiber tracts, the remaining chapters deal with the molecular organization of glutamate synapses assessed by analyses of "prototypical" synapses in the central and peripheral nervous system, or inferred from studies of the regional distribution of specific receptor subtypes or other synaptic proteins. The aim of this volume is to provide an updated account of the chemical anatomy and regional heterogeneity of glutamate synapses. Emphasis has been placed on those aspects that are crucial for an understanding of how signal transmission occurs and of how this process can be modulated in conditions of synaptic plasticity. Thus our intention has been to discuss chemical and structural correlates of the synthesis, synaptic handling, and receptor action of glutamate. Specifically, Chapter 1 focuses on the biochemical compartmentation of glutamate synapses, pathways for glutamate synthesis, and mechanisms of release. Chapter 2 poses the question whether aspartate could act as a cotransmitter with glutamate in certain populations of synapses. Metabotropic and ionotropic glutamate receptors are dealt with in Chapters 3-5, whereas Chapter 6 is concerned with the supramolecular complexes that engage glutamate receptors as well as molecules that are involved in their anchoring and signal transduction. In Chapter 7 the attention is directed to the enzymes that are responsible for the synthesis and degradation of glutamate, and Chapter 8 provides a survey of the expression and functional properties of glutamate transporters. Chapter 9 describes the molecular organization of a peripheral glutamate synapse the first synapse in the auditory system and shows that this synapse shares many of the features of central glutamate synapses, in spite of its distinct embryological origin. The final chapter attempts to correlate chemical, structural, and functional properties of glutamate synapses by using a model synapse that is easily accessible ix

to experimental manipulation u the synapses of the giant reticulospinal axons of the lamprey spinal cord. This chapter is a fitting conclusion of a volume whose task it is to portray a rapidly developing research field where we are now beginning to see how the "chemical anatomy" can be interpreted in terms of the functional demands and physiological properties of the synapse. In 1984, glutamate was the neglected cousin of more well established signaling molecules such as GABA and the monoamines. The dedication of an entire volume of the Handbook to glutamate attests to the fact that 16 years later, glutamate has reached center stage. Oslo, June 2000 OLE PETTER OTTERSEN

JON STORM-MATHISEN

Contents List of Contributors

v

ix

Preface

BIOCHEMISTRY AND ANATOMY OF TRANSMITTER GLUTAMATEJ. BROMAN, B. HASSEL, E. RINVIK AND O.P. OTTERSEN 1. 2.

3.

4.

Introduction Biochemistry of transmitter glutamate 2.1. Synthesis of neuronal glutamate from glucose: some goes via astrocytic lactate 2.2. Glutamine is an important precursor for transmitter glutamate 2.3. Neurons can also carboxylate pyruvate and are therefore not completely dependent on glutamine as a precursor for transmitter glutamate 2.4. Vesicular uptake of transmitter glutamate 2.5. Handling of transmitter glutamate after release: formation of glutamine or pyruvate 2.6. The energy aspect of transmitter glutamate turnover 2.7. Summary Anatomical systems 3.1. Is glutamate immunolabeling evidence of a neurotransmitter role for glutamate? 3.2. Spinal cord 3.2.1. Primary afferent terminals 3.2.2. Intrinsic neurons 3.2.3. Descending inputs 3.2.4. Glutamatergic input to defined spinal neurons 3.2.5. The spinocervical tract 3.3. Brainstem 3.3.1. Medulla oblongata and ports 3.3.2. Midbrain 3.4. Cerebellum 3.5. Thalamus 3.5.1. Corticothalamic projections 3.5.2. Principal subcortical afferents 3.6. Hypothalamus 3.7. Basal ganglia 3.8. Retina 3.9. Cerebral cortex References

1 3 3 5 7 8 8 10 11 11 11 13 13 14 15 16 17 17 17 19 20 23 23 23 24 25 27 28 30

xi

II.

ASPARTATE NEUROCHEMICAL EVIDENCE FOR A TRANSMITTER R O L E - V. GUNDERSEN AND J. STORM-MATHISEN 1. 2. 3.

4. 5. 6. 7.

8. III.

45 45 47 49 50 50 50 51 51 52 53 54 54 55 55 56 56 57 57

METABOTROPIC GLUTAMATE RECEPTORS IMMUNOCYTOCHEMICAL AND IN SITU HYBRIDIZATION ANALYSES- R. SHIGEMOTO AND N. MIZUNO o

2.

xii

Introduction Is aspartate localized in nerve terminals? Is aspartate released by exocytosis from nerve endings? 3.1. Release from synaptosomes 3.2. Release from brain slices 3.3. Release from the intact brain 3.4. Release by heteroexchange? 3.5. Immunocytochemical observations Is aspartate localized in synaptic vesicles? Is aspartate released from a separate pool of nerve endings? The role of the released aspartate Putative aspartatergic neuronal pathways 7.1. The hippocampal formation 7.2. Striatum 7.3. Cerebellar cortex 7.4. Spinal cord 7.5. Auditive systems 7.6. Visual systems References

Introduction Regional and cellular localization of metabotropic glutamate receptors 2.1. An overview 2.2. Distribution of mRNA and immunoreactivity for group I metabotropic glutamate receptors 2.2.1. mGluR1 mRNA 2.2.2. mGluR1 immunoreactivity 2.2.3. mGluR5 mRNA 2.2.4. mGluR5 immunoreactivity 2.3. Distribution of mRNA and immunoreactivity for group II metabotropic glutamate receptors 2.3.1. mGluR2 mRNA 2.3.2. mGluR3 mRNA 2.3.3. mGluR2/3 immunoreactivity 2.3.4. mGluR2 immunoreactivity 2.3.5. mGluR3 immunoreactivity 2.4. Distribution of mRNA and immunoreactivity for group Ill metabotropic glutamate receptors 2.4.1. mGluR4 mRNA 2.4.2. mGluR4 immunoreactivity 2.4.3. Distribution of mRNA and immunoreactivity for mGluR6

63 65 65 76 76 77 78 79 80 80 80 81 82 82 83 83 83 84

3.

4. 5. 6. IV.

2.4.4. mGluR7 mRNA 2.4.5. mGluR7 immunoreactivity 2.4.6. mGluR8 mRNA 2.4.7. mGluR8 immunoreactivity Differential subcellular localization of metabotropic glutamate receptors in relation to transmitter release sites 3.1. mGluRs in postsynaptic elements 3.2. mGluRs in presynaptic elements 3.3. Target-cell-specific segregation of group III mGluRs Abbreviations Acknowledgements References

84 85 86 86 87 87 88 89 90 91 91

AMPA, KAINATE AND NMDA IONOTROPIC GLUTAMATE RECEPTOR EXPRESSION AN IN SITU HYBRIDIZATION ATLAS - W. WISDEN, RH. SEEBURG AND H. MONYER 1. 2.

3.

4. 5.

6.

7.

Introduction AMPA and kainate receptors 2.1. AMPA receptor subunits - - summary of mRNA distribution 2.2. Kainate and 3 receptor subunits - - summary of mRNA distribution NMDA receptors 3.1. NMDA receptor subunits m summary of mRNA distribution 3.1.1. NR 1 RNA splice variants 3.1.2. The NR2 subunits 3.1.3. The NR3A subunit RNA editing Retina 5.1. NMDA receptor subunit mRNAs in the retina 5.2. AMPA receptor subunit mRNAs in the retina 5.3. Kainate receptor subunit mRNAs in the retina Neocortex 6.1. NMDA receptor subunit mRNAs in the neocortex 6.2. NMDA receptor subunit mRNAs in neocortical interneurons 6.3. NR3A expression in neocortex 6.4. AMPA receptor subunit mRNAs in the neocortex 6.5. AMPA receptor subunit mRNAs in neocortical interneurons 6.6. Summary 6.7. Kainate receptor subunit mRNAs in the neocortex Hippocampus 7.1. Hippocampal NMDA receptors 7.1.1. NMDA receptor gene expression in hippocampal principal cells 7.1.2. NMDA receptor subunit gene expression in GABAergic interneurons 7.2. Hippocampal AMPA receptors 7.2.1. AMPA receptor subunit gene expression in hippocampal principal cells

99 99 101 101 104 106 107 109 110 111 111 111 112 113 113 113 114 115 115 116 118 118 119 119 119 121 121 121 xiii

7.2.1.1.

Flip and flop RNA splicing in hippocampal principal cells 7.2.1.2. Development of AMPA receptor flip and flop RNA splicing in hippocampal principal cells 7.2.2. AMPA receptor subunit mRNA in hippocampal intemeurons 7.3. Kainate receptors and ~ subunit in the hippocampus 7.3.1. Kainate receptor subunit mRNA expression in hippocampal principal cells 7.3.2. Kainate receptor subunit mRNA expression in hippocampal interneurons 8. Caudate putamen 8.1. NMDA receptor subunit mRNA distribution in the caudate putamen 8.1.1. NR1 splice variants 8.1.2. NR2 subunit expression 8.1.3. Summary 8.2. AMPA receptor subunit mRNA distribution in the caudate putamen 8.3. Kainate receptor mRNA distribution in the caudate putamen 9. Cerebellum 9.1. NMDA receptor subunit mRNAs in the cerebellum 9.1.1. Purkinje cells 9.1.2. Bergmann glial cells 9.1.3. Granule cells 9.1.4. GABAergic interneurons 9.1.5. Cerebellar nuclei 9.2. AMPA receptor subunit mRNAs in the cerebellum 9.2.1. Purkinje cells 9.2.2. Bergmann glial cells 9.2.3. Granule cells 9.2.4. GABAergic intemeurons 9.2.5. Cerebellar nuclei (medial, interposed and lateral) 9.3. Kainate receptor and 3 subunit mRNAs in the cerebellum 9.3.1. Purkinje cells 9.3.2. Granule cells , 9.3.3. GABAergic intemeurons 10. Spinal cord 10.1. NMDA receptor subunit mRNAs in the lumbar spinal cord 10.2. AMPA receptor subunit mRNAs in the lumbar spinal cord 10.2.1. Dorsal horn 10.2.2. Ventral.horn motor neurons 10.3. Kainate and 3 receptor subunit mRNAs in the spinal cord 11. Acknowledgements 12. References V.

122 122 125 125 126 126 127 128 128 128 129 129 129 130 130 131 131 131 132 132 132 132 132 132 133 133 133 133 133 133 134 135 135 136 137 137 137

REGIONAL AND SYNAPTIC EXPRESSION OF IONOTROPIC GLUTAMATE RECEPTORS- R.S. PETRALIA, M.E. RUB IO, Y.-X. WANG AND R.J. WENTHOLD 1.

xiv

122

Introduction

145

2.

3.

4. 5. VI.

Regional distribution 2.1. Forebrain 2.2. Mid/hindbrain 2.3. Spinal cord and peripheral 2.4. Retina Neuronal distribution 3.1. Synaptic distribution 3.1.1. Adult synapses 3.1.1.1. Differential distribution 3.1.1.2. Tangential distribution 3.1.1.3. Synaptic zones 3.1.2. Developing synapses 3.2. Cytoplasmic distribution 3.3. Functional considerations 3.3.1. Targeting mechanisms 3.3.2. Insertion and removal of receptors at the synapse Distribution in glia References

145 149 153 155 157 158 158 158 158 162 162 165 168 169 169 172 173 174

TARGETING AND ANCHORING OF GLUTAMATE RECEPTORS AND ASSOCIATED SIGNALING M O L E C U L E S - M. WYSZYNSKI AND M. SHENG 1. 2.

3.

4. 5. 6. 7. 8.

Introduction NMDA receptors 2.1. Association of NMDA receptors with the PSD 2.2. Interactions of the NR2 subunit: the PSD-95 complex 2.3. Synaptic targeting by PSD-95 2.4. Assembly of a signaling complex by PSD-95 2.5. Anchoring to the cytoskeleton via PSD-95 2.6. Interactions of the NR1 subunit 2.7. Other interactions of NMDA receptors AMPA receptors 3.1. Synaptic targeting of AMPA receptors 3.2. Interactions with PDZ proteins 3.3. Interactions with NSF and signaling proteins Kainate receptors and ~ receptors Metabotropic glutamate receptors Concluding comments: comparing glutamate receptors Acknowledgements References

183 183 183 184 185 186 188 189 190 190 190 191 192 193 193 195 196 197

VII. ENZYMES RESPONSIBLE FOR GLUTAMATE SYNTHESIS AND DEGRADATION- T. KANEKO 1. 2.

Introduction Distribution of glutaminase in the nervous system 2.1. Forebrain regions

203 204 205 XV

o

4. 5. 6. 7.

2.2. Diencephalic regions 2.3. Brainstem and cerebellar regions 2.4. Spinal cord and peripheral nerves 2.5. Retina 2.6. Non-neural distribution of glutaminase Glutamate synthesis and metabolism in glial cells Glutamate and AAT in GABA synthesis Concluding remarks Acknowledgements References

211 215 217 218 218 219 221 225 227 227

VIII. SODIUM- AND POTASSIUM-DEPENDENT EXCITATORY AMINO ACID TRANSPORTERS IN BRAIN PLASMA MEMBRANES - N.C. DANBOLT 1. 2. 3. 4.

Introduction Glutamate transporter types Mechanism of glutamate uptake Localization of glutamate transporters 4.1. Localization of GLT (EAAT2) 4.1.1. GLT is the major glutamate transporter in the forebrain 4.1.2. Exclusive glial expression of GLT protein, but not of GLT mRNA 4.1.3. GLT protein in neurons 4.1.4. Regional and subcellular distribution of GLT in adult rat brain tissue 4.2. Localization of GLAST (EAAT1) 4.2.1. Cellular distribution of GLAST in the CNS 4.2.2. Subcellular distribution of GLAST 4.2.3. Concentrations of GLAST protein 4.3. Localization of EAAC (EAAT3) 4.3.1. Antibodies to EAAC 4.3.2. Localization of EAAC in the adult CNS 4.4. Localization of EAAT4 4.4.1. Regional and cellular distribution of EAAT4 4.4.2. Subcellular distribution in the adult Purkinje cells 4.5. Localization of EAAT5 4.6. Developmental changes in glutamate transporter expressions 4.6.1. Changes in transporter concentrations 4.6.2. Changes in the localizations of GLT and GLAST 5. Regulation of glutamate uptake 5.1. Glutamate transporter expression 5.2. Posttranslational regulation of transporters 6. The role of glutamate uptake in synaptic transmission 6.1. Overview 6.2. The time course of glutamate in the synaptic cleft 6.3. Densities of glutamate transporters and paradoxical effects 6.4. Intersynaptic crosstalk 7. Concluding remarks

xvi

231 232 232 233 233 233 234 235 235 236 236 237 237 238 238 238 239 239 239 240 240 240 241 241 241 242 243 243 243 244 244 245

8. Abbreviations 9. Acknowledgements 10. References IX.

GLUTAMATE NEUROTRANSMISSION IN THE MAMMALIAN INNER EAR - S. USAMI, A. MATSUBARA, S. FUJITA, Y. TAKUMI AND O.R OTTERSEN 1. 2. 3.

Introduction Glutamate in hair cells A glutamate-glutamine cycle in the inner ear? Glutamine synthetase and glutamate transporters 4. Distribution of phosphate-activated glutaminase in the inner ear 5. Glutamate release 6. Glutamate receptors 6.1. AMPA receptors 6.2. Other types of glutamate receptor 7. Pathology of the glutamatergic synapse 8. Conclusion 9. Acknowledgements 10. References Xo

246 246 246

255 255 258 260 262 262 262 266 266 267 268 268

A MODEL GLUTAMATE SYNAPSE - - THE LAMPREY GIANT RETICULOSPINAL A X O N - O. SHUPLIAKOV AND L. BRODIN 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11.

Introduction The lamprey reticulospinal synapse - - an overview Organization of the reticulospinal axon Synaptic localization of glutamate and related amino acids Synaptic vesicle pools Presynaptic Ca 2+ channels Presynaptic modulation of transmitter release Synaptic vesicle recycling Molecular mechanisms in synaptic vesicle endocytosis Conclusions References

Subject Index

273 273 274 276 279 279 281 284 286 286 287

289

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CHAPTER I

Biochemistry and anatomy of transmitter glutamate J. BROMAN, B. HASSEL, E. RINVIK AND O.E OTTERSEN

1. INTRODUCTION The powerful excitatory effect of glutamate (Glu) on central neurons was discovered more than forty years ago (Hayashi, 1954; Curtis and Watkins, 1960). However, as Glu is present in high concentrations and is relatively evenly distributed among different brain areas, it took a long time until Glu was generally accepted as a neurotransmitter (see Krnjevic, 1986; Watkins, 1986). By the mid-1980s, Glu largely fulfilled the four main criteria for classification as a neurotransmitter, i.e.: (1) presynaptic localization; (2) release by physiological stimuli; (3) identical action with naturally occurring transmitter; and (4) mechanism for rapid termination of transmitter action (Fonnum, 1984). Later investigations have strengthened a neurotransmitter role for Glu. Such investigations include the demonstration of ATP-dependent selective transport of Glu into purified synaptic vesicles (Naito and Ueda, 1985; Maycox et al., 1988; Fykse et al., 1989; Winter and Ueda, 1993), the presence of high concentrations of Glu in synaptic vesicles isolated from the brain (Riveros et al., 1986; Burger et al., 1989; Orrego and Villanueva, 1993), and a Ca2+-dependent exocytotic release of Glu from isolated nerve terminals (Nicholls, 1995). Rapid application of Glu to neuronal membrane patches at a concentration (1 raM) similar to that estimated to be present in the synaptic cleft following exocytotic release, mimics the postsynaptic response following activation of excitatory synapses (Clements et al., 1992; Colquhoun et al., 1992; Bergles et al., 1999). Extensive molecular studies during the recent decade have also provided detailed knowledge on the subunit proteins and gene families of Glu receptors (Anwyl, 1995; Blackstone and Huganir, 1995), the distribution of which has been mapped by in situ hybridization and immunocytochemistry (see Chapters 3-6). Glutamate has now gained an indisputable neurotransmitter status and has been localized to a large number of fiber systems (Figs. 3-7). But other endogenous excitatory amino acids have also been suggested to act as transmitters. The evidence supporting a neurotransmitter role of aspartate the most prevalent endogenous excitatory amino acid after Glu is reviewed in Chapter 2. Many different approaches have been used to identify the neurons that use Glu as a transmitter. Biochemical techniques, including analysis of reduced content or uptake of Glu or Glu analogues following lesions, have proved useful in investigations of major projections (e.g. corticofugal fiber tracts; Fonnum, 1984; Storm-Mathisen and Ottersen, 1988; Ottersen, 1991), but poor sensitivity hampers analyses of less massive pathways. Detection of many minor glutamatergic projections was made possible by the use of the metabolically inert Glu Handbook of Chemical Neuroanatom~; Vol. 18: Glutamate O.P. Ottersen and J. Storm-Mathisen, editors (~ 2000 Elsevier Science B.V. All rights reserved.

Ch. I

J. Broman et al.

analogue D-[3H]aspartate as a transmitter-specific retrograde tracer (Baughman and Gilbert, 1980; Streit, 1980; Ottersen, 1991). However, D-[3H]aspartate does not differentiate between putative glutamatergic and aspartergic projections. There are also a number of fiber tracts likely to use Glu as a neurotransmitter that are poorly labeled or unlabeled by D-[3H]aspartate, possibly due to low presynaptic Glu uptake capacity of the terminals of such pathways (Ottersen, 1991). To delineate glutamatergic pathways in the CNS, alternative methods were needed that could unravel the detailed anatomical distribution of Glu. A tool for microscopical demonstration of Glu came with the introduction of amino acid immunocytochemistry (Storm-Mathisen et al., 1983). Antibodies raised against aldehyde-fixed Glu and GABA were used to generate a map of the distribution of the respective amino acids that was published in an early volume of this Handbook Series (Ottersen and Storm-Mathisen, 1984a). Soon several other groups raised antisera to amino acids and used these antisera for visualizing amino acids in the brain and spinal cord (Hodgson et al., 1985; Wanaka et al., 1987; Yoshida et al., 1987; Hepler et al., 1988; Chagnaud et al., 1989; Liu et al., 1989; Pow and Crook, 1993). In accordance with biochemical data, immunocytochemical studies demonstrated that Glu is widely distributed in the brain and localized not only in presumed glutamatergic neurons but also in neurons with other transmitter signatures. This was not surprising, taking into account the involvement of Glu in several metabolic functions (protein synthesis, intermediary metabolism, and as a precursor for GABA). The ubiquity of Glu, and the inability of Glu antisera to differentiate between metabolic and transmitter pools, called for a quantitative approach that could be applied to the nerve terminals. The post-embedding immunogold technique (Figs. 3 and 6) was shown to meet these demands (Somogyi and Hodgson, 1985; Somogyi et al., 1986). The interpretation of immunogold data for Glu or other antigens requires knowledge of the degree of labeling specificity and of the relationship between labeling density and antigen concentration. Using model systems that were designed to address these questions (Fig. 6D; Ottersen, 1987, 1989) it was demonstrated that a close to linear relationship between gold particle density and concentration of fixed Glu can be achieved within the biological relevant range of Glu concentrations. To examine Glu content in terminals that cannot be identified solely by morphological criteria, combinations of anterograde tracing and immunogold labeling have been developed (De Biasi and Rustioni, 1988; Broman et al., 1990). Quantitative analysis of Glu immunogold-labeled preparations has become a widely used and fruitful tool in the identification of putative glutamatergic nerve terminals. As indicated above, Glu is not only a neurotransmitter but is also involved in a variety of metabolic functions in the brain. The metabolism of Glu is complicated and involves neurons as well as glial cells. Transmitter Glu may be synthesized through different metabolic pathways, and different populations of glutamatergic neurons may differ in certain aspects of Glu metabolism. The first part of this chapter will provide an update on the metabolism of Glu and related compounds in the brain. The second part will deal with anatomical aspects of transmitter Glu and provide an overview of the neuronal populations that use Glu as a neurotransmitter. As Glu immunogold data have not been reviewed in this Handbook Series (except in chapters on specific regions, e.g. Jones, 1998) we will devote much of Section 3 to these. Reference to earlier work with other techniques will largely be made through citation of review articles (e.g. Ottersen and Storm-Mathisen, 1984a; Fonnum, 1984, 1991; Storm-Mathisen and Ottersen, 1988; Ottersen, 1991; Fonnum and Hassel, 1995; Storm-Mathisen et al., 1995). The reader is referred to these publications for a complete bibliography.

Biochemistry and anatomy of transmitter glutamate

Ch. I

2. BIOCHEMISTRY OF TRANSMITTER GLUTAMATE

The formation and degradation of Glu is a part of the general energy metabolism of the brain, since glucose, which is the main, possibly the only, physiological energy substrate for the brain, is converted almost stoichiometrically into Glu before being oxidized further via the tricarboxylic acid (TCA) cycle. Because all brain cells contain Glu as a byproduct of energy metabolism, a neuron can be defined as glutamatergic on an immunocytochemical basis only after detection of Glu in synaptic vesicles; the presence of Glu in neuronal cell bodies is of little or no value for the determination of neurotransmitter identity. In the brain, Glu is present in separate pools. It is customary to refer to the transmitter pool (located in vesicles of glutamatergic terminals), the pool of Glu that serves as precursor of GABA (located in GABAergic neurons), the pool of Glu that serves as precursor of glutamine (located in glia), and lastly the metabolic pool of Glu (present in all cells) which is a byproduct of energy metabolism. The various pools communicate with each other, for instance when Glu is diverted from the metabolic pool to become transmitter or precursor of GABA and glutamine, and when the amino acid transmitters return to the metabolic pool and are metabolized to CO2 and water. Further, there is extensive transport of Glu and its derivatives, GABA and glutamine, between cell types. In the following we will discuss the formation of transmitter Glu, its storage in synaptic vesicles, the inactivation of transmitter Glu by uptake into astrocytes and conversion to non-transmitter metabolites. Finally, we will estimate the energy cost of glutamatergic neurotransmission. 2.1. SYNTHESIS OF NEURONAL GLUTAMATE FROM GLUCOSE: SOME GOES VIA ASTROCYTIC LACTATE

Serum glucose is by far the most important precursor for transmitter Glu, since of the various possible Glu precursors present in serum, only glucose shows a consistent arteriovenous difference (Gibbs et al., 1942). Glucose transport into the brain has a Km of 6-9 mM, consistent with the normal serum level of glucose. Glucose enters the brain by crossing the blood-brain barrier and the astrocytic interphase constituted by the perivascular end feet surrounding brain capillaries. The uptake is mediated by a specific transporter, GLUT1 (Maher, 1995; Morgello et al., 1995), that is expressed by both endothelial cells and astrocytes. In recent years it has become clear that some of the glucose that enters the brain is metabolized glycolytically by astrocytes to lactate which in turn is given off to the extracellular fluid and taken up by neurons (Brazi~ikos and Tsacopoulos, 1991; for review, see Tsacopoulos and Magistretti, 1996). This view is supported by recent findings that the extracellular concentration of glucose in the brain in the awake rat is quite low: 0.2-1 mM (Lowry et al., 1998; McNay and Gold, 1999). If we assume that glucose enters neurons only from the extracellular fluid after having passed through astrocytes, then 85-95% of the serum glucose that enters the brain must be metabolized glycolytically by astrocytes. Because the cerebral glucose transporters are facilitative and sodium-independent (e.g. Asano et al., 1992), it follows that for a glucose gradient to be present over the neuronal cell membrane, the intraneuronal concentration of glucose must be very low. However, some findings point to glucose as such as a quantitatively important energy substrate for neurons. First, the regional uptake of the glucose analogue, 2-deoxyglucose, matches the regional expression of the neuronal glucose transporter, GLUT3, not that of the glial GLUT1 (Maher et al., 1994). Second, glycolytic enzymes are highly expressed in

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neurons in vivo, apparently more so than in astrocytes (e.g. Oster-Granite and Gearhart, 1980; Zeitschel et al., 1996; Cimino et al., 1998). Third, cultured neurons metabolize glucose more avidly than do cultured astrocytes (e.g. Olsen et al., 1999). The low extracellular concentration of glucose seems to correspond well with the low Km for glucose found in synaptosomal preparations, 0.2-0.3 mM (Diamond and Fishman, 1973; Heaton and Bachelard, 1973), but the low Km may reflect the hexokinase activity of synaptosomes (Kin -- 50 I~M) (Maher et al., 1996); hexokinase, which catalyzes phosphorylation of glucose (or 2-deoxyglucose), is generally thought to control the influx of glucose into the brain (Whitesell et al., 1995). When expressed in hexokinase-poor Chinese hamster oocytes, GLUT3 has a Km for glucose of 2-3 mM (Asano et al., 1992; Maher et al., 1996). At present, therefore, we do not know the relative importance of neurons and astrocytes in the initial metabolism of glucose. The serum concentration of lactate is 1-3 mM, and the extracellular concentration in the brain is 0.2-0.4 mM (Herrera-Marschitz et al., 1996; Demestre et al., 1997). Therefore, astrocytes, which take up serum lactate, probably act as a lactate reservoir, buffering the extracellular concentration of lactate. The anxiogenic effect of high levels of serum lactate (Pitts and McClure, 1967; Dager et al., 1997) may reflect the need for such buffering. Lactate is taken up by monocarboxylate/H + co-transporters (Broer et al., 1999a) along the lactate gradient and the intraneuronal concentration of lactate must therefore be lower than that of the extracellular fluid. Lactate is avidly metabolized by neurons in vivo, but hardly at all by astrocytes (O'Neal and Koeppe, 1966; Hassel and Br~the, 2000a). In neurons, lactate is converted to pyruvate and hence to acetyl-coenzyme A which condenses with oxaloacetate to form citrate. Citrate, in turn, is converted to isocitrate and hence to ~-ketoglutarate from which Glu is formed (Fig. 1). The time scale of these reactions is illustrated by the strong labeling of neuronal Glu 2-5 min after an intravenous bolus injection of isotopically labeled glucose (Van den Berg et al., 1969; Hassel and Sonnewald, 1995a); isotopically labeled lactate leads to even more rapid labeling of neuronal Glu (Hassel and Brfithe, 2000a). The cerebral TCA cycle activity is 15-20 nmol min -1 mg -1 protein (Gaitonde, 1965; Borgstr6m et al., 1976; Sokoloff et al., 1977; Lu et al., 1983; Mason et al., 1992, 1995). This activity corresponds quite well to the whole brain activity of ~-ketoglutarate dehydrogenase, and it is lower than

~In astrocytes

Anaplerosis: Fig. 1.

~ ln GABAergicneurons

Simplified scheme of the TCA cycle and the formation of glutamate from ~-ketoglutarate (~-kg). In astrocytes glutamate is amidated to glutamine; in GABAergic neurons some of the glutamate is decarboxylated and enters the GABA shunt. In both neurons and astrocytes anaplerosis occurs via carboxylation of pyruvate to malate or oxaloacetate (ox-ac) from which aspartate is formed.

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all other enzyme activities of the TCA cycle as measured in vitro. Therefore, ot-ketoglutarate dehydrogenase, which converts c~-ketoglutarate into succinyl-CoA, is a rate-limiting step of the TCA cycle (Lai et al., 1977), a bottleneck that causes c~-ketoglutarate to build up. ~-Ketoglutarate is transaminated to Glu by the highly active transaminases, especially aspartate aminotransferase (cf. Mason et al., 1992) which uses aspartate as an amino group donor, and alanine aminotransferase, which uses alanine as the amino group donor. Alanine is exported from astrocytes and taken up by neurons (Sonnewald et al., 1991; Westergaard et al., 1993). Accordingly, alanine injected into rat striatum in vivo is taken up by neurons and metabolized to Glu (Fonnum et al., 1997). Other possible amino group donors are the branched chain amino acids, especially leucine, which enters the brain from the circulation (Yudkoff, 1997). The large pool of Glu present in glutamatergic neurons is therefore maintained by the bottleneck function of et-ketoglutarate dehydrogenase in the TCA cycle, the very high activities of the transaminases compared to et-ketoglutarate dehydrogenase, and by the ample supply of amino group donors in transamination reactions. The low level of Glu in GABAergic neurons and in astrocytes (Fig. 6) is probably due to the fact that the bottleneck of ct-ketoglutarate dehydrogenase is bypassed in these cell types. In GABAergic neurons Glu enters the GABA shunt and is converted successively into GABA, succinic semialdehyde and succinyl-CoA. This pathway is parallel to the 0L-ketoglutarate dehydrogenase reaction, and in awake mice it has been calculated that the fluxes through the GABA shunt and the ~-ketoglutarate dehydrogenase reaction are fairly similar (Hassel et al., 1998). This is probably also the reason why the level of aspartate is high in the cell bodies of GABAergic neurons (Ottersen and Storm-Mathisen, 1985; Hassel et al., 1992, 1995a; Hassel and Sonnewald, 1995b): the citrate synthase reaction is limited by the availability of acetylCoA which is provided by pyruvate dehydrogenase (Lai et al., 1977). Therefore, oxaloacetate may build up in GABAergic neurons, leading to formation of a large pool of aspartate (cf. Fig. 1) in the same way that build-up of et-ketoglutarate in glutamatergic neurons leads to accumulation of Glu. In astrocytes Glu is diverted from the bottleneck of ot-ketoglutarate dehydrogenase by the formation of glutamine which leaves the cells. Accordingly, the levels of both Glu and aspartate are low in astrocytes (Ottersen and Storm-Mathisen, 1985). 2.2. GLUTAMINE IS AN IMPORTANT PRECURSOR FOR TRANSMITTER GLUTAMATE Although the above section describes the formation of Glu in neurons, it has been assumed by many researchers that glutamine is the main, maybe the only, immediate precursor for transmitter Glu. Glutamine is formed from Glu by amidation; in the brain the glutaminesynthesizing enzyme, glutamine synthetase, has a strictly astrocytic and oligodendroglial localization (Martinez-Hernandez et al., 1977; Tansey et al., 1991; Miyake and Kitamura, 1992). It has been calculated that ~60% of the 0t-ketoglutarate formed in astrocytes is converted to Glu and hence to glutamine both in vitro and in vivo (Hassel et al., 1994, 1995b). Because astrocytes in vivo do not express glutaminase (Akiyama et al., 1990; Ottersen et al., 1998; Laake et al., 1999), the enzyme which converts glutamine into Glu, it may be assumed that most of the glutamine formed in glia is exported to the extracellular fluid where the concentration is quite high, 0.2 mM (Lerma et al., 1986). Two glutamine carriers that could regulate the efflux of glutamine from astrocytes have recently been identified (Broer et al., 1999b; Chaudhry et al., 1999). In cultured neurons three different glutamine carriers that mediate glutamine uptake have been identified (Tamarappoo et al., 1997), but so far they have not been cloned, and the

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distribution in the brain has not yet been established. The uptake of glutamine into nerve terminals occurs against a concentration gradient, since the extracellular concentration is ~0.2 mM, whereas the intracellular concentration may be up to several millimolars (Ottersen et al., 1992; also see Fig. 6A-C). Exogenous glutamine has been found to be a good precursor for releasable Glu in vitro (Cotman and Hamberger, 1978; Hamberger et al., 1979; Reubi, 1980; Ward et al., 1983), but because glutaminase is strongly inhibited by its products, Glu and ammonia, which may become diluted by buffers in the in vitro setting, the enzyme activity may easily be overestimated in vitro (Fonnum, 1993). Another source of in vitro artifacts which applies to cultured brain cells is the common use of culture media with a high concentration of glutamine, 2-2.5 mM. The continuous exposure to such concentrations, which are ten times that of the extracellular fluid in the brain, could induce glutamine dependence. As pointed out by Fonnum (1991), the precursor role of glutamine has been difficult to demonstrate in vivo with the use of radiolabeled glutamine, although many neuronal populations express glutaminase (Donoghue et al., 1985; Akiyama et al., 1990; Ottersen et al., 1998; Laake et al., 1999). Radiolabeled, i.e. exogenous, glutamine has had to be administered in large amounts to intact brain tissue and over surprisingly long time periods to achieve radiolabeling of releasable transmitter amino acids (Thanki et al., 1983). As shown by Zielke et al. (1998), glutamine injected intracerebrally is to a large extent metabolized to CO2 and water, which agrees with the role of glutamine as an energy source for neurons (Bradford et al., 1978; Hassel et al., 1995b). The high extracellular level of glutamine in the brain, which dilutes the injected radiolabeled glutamine, does not explain the low labeling of transmitter Glu, since intracerebral injection of radiolabeled glucose labels Glu very efficiently (e.g. Hassel et al., 1992) in spite of a high level of extracellular glucose: in anesthetized animals extracellular glucose may reach 3 mM (Ronne-Engstrom et al., 1995). One may speculate whether exogenous and endogenous glutamine are handled differently by the brain. To study the metabolic fate of endogenous glutamine one can use isotopically labeled substrates that are taken up selectively by astrocytes, such as acetate, propionate or butyrate. Intracerebral or intravenous injection of isotopically labeled acetate leads to strong labeling of endogenous glutamine and, after export to neurons, to labeling of neuronal Glu and GABA (O'Neal and Koeppe, 1966; Van den Berg et al., 1966, 1969; Cerdan et al., 1990; Chapa et al., 1995; Hassel et al., 1995b, 1997). Inhibition of synthesis of (endogenous) glutamine in vivo with methionine sulfoximine, an inhibitor of glutamine synthetase, or fluorocitrate, an inhibitor of the astrocytic TCA cycle, reduces the release of transmitter Glu and GABA as determined by microdialysis (Paulsen et al., 1988; Paulsen and Fonnum, 1989). These results, although obtained by indirect methods, do support the idea of glutamine as an important precursor for transmitter Glu in vivo. Glutaminase is located on the external aspect of the inner mitochondrial membrane (Roberg et al., 1995; Fig. 6E). Such a localization could suggest that the Glu which is formed from glutamine is largely returned to the cytosol without first equilibrating with intramitochondrial Glu, meaning that the transmitter pool of Glu (i.e. that derived from glutamine) is different from the metabolic pool of Glu. However, because glutamine is an important energy substrate for neurons (Bradford et al., 1978; Hassel et al., 1995b), much of the Glu that is formed from glutamine must enter mitochondria. Glutaminase may become of special importance after cell damage, e.g. as caused by trauma or hypoxia, when the enzyme leaks out of neurons and into the extracellular space. Here it may convert extracellular glutamine into Glu, thus contributing to a continuous and excitotoxic glutamatergic stimulation of neurons. Such a mechanism has been demonstrated

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in vitro (Driscoll et al., 1993; Newcomb et al., 1997), and may also be operative in vivo (Newcomb et al., 1998). Astrocytic export of glutamine implies a continuous loss of ~-ketoglutarate from the astrocytic TCA cycle. This loss has to be compensated, otherwise the astrocytic TCA cycle would be drained of its intermediates, and the ability to generate ATP would be impaired. In the brain, with its restricted entry of TCA cycle intermediates (e.g. citrate) across the blood-brain barrier, the only way to replenish such a loss is through the anaplerotic process of pyruvate carboxylation, by which pyruvate (derived from glucose via glycolysis) receives a carboxylic group in the form of CO2 and is converted to oxaloacetate or malate (Figs. 1 and 2). In vivo and in vitro it has been shown that astrocytic pyruvate carboxylation corresponds quite closely to the formation of glutamine (Hassel et al., 1995b; Gamberino et al., 1997). Astrocytes express the enzymes pyruvate carboxylase (Yu et al., 1983; Shank et al., 1985; Cesar and Hamprecht, 1995) and cytosolic and mitochondrial malic enzyme (Kurz et al., 1993; McKenna et al., 1995), the three pyruvate-carboxylating enzymes in brain (Salganicoff and Koeppe, 1968). 2.3. NEURONS CAN ALSO CARBOXYLATE PYRUVATE AND ARE THEREFORE NOT COMPLETELY DEPENDENT ON GLUTAMINE AS A PRECURSOR FOR TRANSMITTER GLUTAMATE Glutamatergic neurotransmission implies a loss of Glu from glutamatergic neurons, because transmitter Glu to a large extent is taken up by astrocytes. A net loss of Glu implies a loss of ~-ketoglutarate from the neuronal TCA cycle that would cause a reduction in ATP production. Anaplerosis, i.e. carboxylation of pyruvate to malate or oxaloacetate (Fig. 1) is therefore required. For many years it has been assumed that astrocytes were the only brain cells capable of pyruvate carboxylation, so that the loss of Glu from neurons would have to be compensated by uptake of glutamine from astrocytes. The main reason for this assumption was the finding of the enzyme pyruvate carboxylase in astrocytes and not in neurons (Yu et al., 1983; Shank et al., 1985). Earlier, Patel (1974) had published a study which suggested that pyruvate carboxylase was by far the most active pyruvate-carboxylating enzyme in the brain. Taken together these studies indicated that astrocytes were the main, perhaps the only, anaplerotic compartment in the brain, a notion which seemingly received support from the observation that intravenous infusion of radiolabeled bicarbonate led to better labeling of glutamine than of Glu (Waelsch et al., 1964). The latter finding was taken to imply that pyruvate carboxylation occurred in the glutamine-synthesizing cells, i.e. glia. These findings formed the basis for the concept of a glutamine cycle (Van den Berg and Garfinkel, 1971; Benjamin and Quastel, 1975), the 1:1 exchange between astrocytes and neurons of glutamine for Glu and GABA. However, in the study of Waelsch et al. (1964) the radiolabeled bicarbonate given intravenously would mainly reach the astrocytic compartment via the astrocytic end feet that envelop brain capillaries. When given intracerebrally, the radiolabel also reaches the neuronal compartment, and Glu is labeled to a greater extent than glutamine (Hassel and Br~the, 2000b). Similarly, cultured neurons show very active pyruvate carboxylation (Hassel and Br~the, 2000b); in this study any contribution from astrocytes that might contaminate the neuronal cultures was avoided by pretreating the cultures with the gliotoxin fluoroacetate. Regarding the enzymatic pathway, malic enzyme activity was recently found in synaptosomes (Cruz et al., 1998) and the mitochondrial isoform was detected by immunohistochemistry in cultured neurons (Vogel et al., 1998). Three decades earlier Salganicoff and Koeppe (1968) showed that the mitochondrial malic enzyme in brain had a high pyruvate carboxylating activity.

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The finding that neurons, or at least subpopulations of neurons, seem to have the ability to replenish their TCA cycle by carboxylating pyruvate (Hassel and Brfithe, 2000a,b) may explain why some glutamatergic pathways have a low level of glutaminase, whereas others have high levels as detected by immunocytochemistry (Laake et al., 1999; Fig. 6E), and it explains how transmitter Glu may be formed from neuronal precursors. 2.4. VESICULAR UPTAKE OF TRANSMITTER GLUTAMATE The uptake of Glu in synaptic vesicles is one of the criteria for the definition of Glu as a neurotransmitter. Based on lesion experiments in which nerve terminals were caused to degenerate, the transmitter pool of Glu has been estimated to be 20-30% of the total brain Glu content (Lund-Karlsen and Fonnum, 1978; Walaas and Fonnum, 1980; Fonnum et al., 1981). The Glu formed in the nerve terminals enters the synaptic vesicles via a transporter that is not yet cloned. The vesicular transporter has a low affinity for Glu, with a Km around 1 mM (Naito and Ueda, 1985; Maycox et al., 1988). This is ~ 1000 times higher than the Km of the plasma membrane transporters, which agrees with the concentration of Glu being 1000-fold higher in the cytosol than in the extracellular fluid. The transport of Glu into vesicles is driven by an electrochemical gradient generated by a proton pump which is dependent on ATP and magnesium and is stimulated by a chloride concentration of 4-10 mM, similar to the cytosolic concentration (Naito and Ueda, 1983, 1985; Maycox et al., 1988; Fykse et al., 1989). The vesicular concentration of Glu has been estimated to ~ 100 mM, which is in good agreement with experimental data (Burger et al., 1989; Shupliakov et al., 1992). Depolarization of glutamatergic neurons leads to influx of calcium into the terminal, which triggers exocytosis of Glu by fusion of the membrane of the synaptic vesicle with the plasma membrane. This fusion is mediated by the interaction of vesicular proteins with plasma membrane proteins, a process which to a large extent is regulated by protein phosphorylation (reviewed by Hanson et al., 1997), and which therefore is ATP-dependent (e.g. Esser et al., 1998). 2.5. HANDLING OF TRANSMITTER GLUTAMATE AFTER RELEASE: FORMATION OF GLUTAMINE OR PYRUVATE After its release transmitter Glu must be cleared from the synaptic cleft. It is a matter of debate whether the plasma membrane transporters located in astrocytic and neuronal cell membranes in the vicinity of the synapse are capable of actually removing the Glu fast enough to account for the rapid clearance of transmitter from the cleft, or whether they act (on a short time scale) by binding Glu (Lehre and Danbolt, 1998). But once internalized into astrocytes, Glu may enter one of two major biochemical pathways (Fig. 2). First, Glu may become amidated to glutamine by glutamine synthetase in the astrocytic cytosol. This glutamine presumably equilibrates with the general pool of astrocytic glutamine. The detection of glutamine synthetase in astrocytic processes in the vicinity of glutamatergic synapses indicates the importance of this pathway (Derouiche and Frotscher, 1991). Second, Glu may enter the mitochondria of astrocytes to become transaminated (by aminotransferases) or deaminated (by glutamate dehydrogenase) to 0~-ketoglutarate and may be oxidized successively to succinate, fumarate and malate. Malate may become decarboxylated to pyruvate, presumably after leaving the mitochondria because the most likely candidate for this decarboxylation is cytosolic malic enzyme, which is strongly expressed by astrocytes (Kurz et al., 1993). In cultured astrocytes it has been shown that the higher the extracellular concentration of Glu the more pyruvate (and hence lactate) will be formed via malate decarboxylation (McKenna

Biochemistry and anatomy of transmitter glutamate

(i

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/

Fig. 2. Metabolic interactions between neurons and astrocytes. Glucose enters the brain through the astrocytic end feet that envelop brain capillaries. In the astrocytes some of the glucose is metabolized to lactate which is exported to the extracellular fluid and taken up by neurons. In neurons lactate is converted to pyruvate which is either decarboxylated to acetyl-CoA or carboxylated to malate to enter the TCA cycle. Glutamate may therefore be formed in neurons from e~-ketoglutarateor from glutamine, which is imported from astrocytes. The glutamate that is released is taken up by astrocytes and amidated to glutamine or metabolized via the TCA cycle. The malate thus formed may leave the TCA cycle and become decarboxylated to pyruvate and lactate. For lack of space, astrocytic pyruvate carboxylation is indicated only by the reversible formation of lactate. Notice that the relative importance of the various pathways in vivo is a matter of debate (see text).

et al., 1996). The lactate thus formed from transmitter Glu is probably also shunted back to neurons, but it has been proposed that it may serve a distinct function as a vasodilator in the brain, coupling glutamatergic neurotransmission to an increase in cerebral blood flow (Hassel and Sonnewald, 1995a): lactate is a vasodilator in the brain, irrespective of pH (Laptook et al., 1988). Malate may of course also be oxidized further in the astrocytic TCA cycle, since malate has been shown to be an excellent substrate for astrocytes (McKenna et al., 1990). The magnitude of the flux of transmitter Glu from neurons to astrocytes may be roughly calculated from the formation of glutamine from transmitter Glu. A problem is that glutamine may be formed not only from transmitter Glu or GABA, but also from o~-ketoglutarate derived from the astrocytic TCA cycle. In a series of papers Shulman, Rothman, Behar, Mason, and colleagues have addressed this issue with the use of 13C nuclear magnetic resonance spectroscopy (NMRS) in combination with i.v. infusion of [1-13C]glucose (Mason et al., 1992, 1995; Sibson et al., 1997, 1998; Shen et al., 1999). The authors base their calculations on the fact that the 13C-labeling of glutamine lags behind the labeling of Glu when [1-13C]glucose is the precursor. This lag is assumed to represent the time needed for 13C-labeled transmitter Glu to reach astrocytes for amidation to glutamine. Given the insensitivity of the 13C NMRS technique, which could underestimate the 13C-labeling of glutamine and overestimate the lag in glutamine labeling, their calculation that 90% of glutamine is formed from transmitter Glu, is probably an overestimation. In another study, the formation of glutamine from transmitter Glu was 40% of the total formation of glutamine (Hassel et al., 1997). This value was determined in mice treated with fluoroacetate, which causes somnolence, and is probably an

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underestimation. In the following we will therefore assume that 50-80% of brain glutamine is formed from transmitter Glu. In the rat and human brain the level of glutamine is 60 nmol/mg protein, of which 50-80%, i.e. 30-50 nmol/mg protein, may be formed from transmitter Glu. This value corresponds to the transmitter Glu pool size (20-30% of a brain level of 100-120 nmol Glu/mg protein). The flux of transmitter Glu to astrocytic glutamine would then be 20-30% of the whole brain turnover rate for Glu (16-20 nmol mg protein -1 min-1), i.e. 3-6 nmol mg protein -1 min-1; this value is similar to the value of 2.1 nmol mg protein -1 min -~ obtained in anesthetized rats (Sibson et al., 1997). Because some of the transmitter Glu may be metabolized via non-glutamine pathways, e.g. to lactate (Hassel and Sonnewald, 1995a; McKenna et al., 1996), the total flux of Glu to astrocytes may be somewhat higher. 2.6. THE ENERGY ASPECT OF TRANSMITTER GLUTAMATE TURNOVER Several of the steps in the formation and degradation of transmitter Glu has a cost in terms of ATP expenditure. Uptake of Glu into vesicles is ATP-dependent. The stoichiometry has not been determined, but extrapolating from the plasma membrane transporter and from the > 100-fold higher concentration of Glu inside the vesicle than in the cytosol, it is likely that one molecule of ATP is consumed per molecule of Glu. Fusion of the vesicular membrane with the plasma membrane depends on protein phosphorylation and is therefore also ATP-dependent. However, since each vesicle contains approximately a thousand molecules of Glu the ATP utilization per molecule of Glu is low. (A vesicular inner radius of 17 nm gives a vesicular volume of 2 x 10 -20 1, a vesicular concentration of 100 mM Glu equals 6 x 10 22 molecules/l; the product is 1200 molecules per vesicle.) Uptake of Glu into astrocytes is coupled to influx of three molecules of sodium (Levy et al., 1998) which are cleared by the Na/K-ATPase, leading to the use of one molecule of ATP per molecule of internalized Glu. Formation of glutamine from Glu requires one ATP per Glu. Even when glutamine is formed from c~-ketoglutarate derived from the astrocytic TCA cycle, this loss is compensated by pyruvate carboxylase activity, using one ATP per molecule of oxaloacetate produced (Scrutton et al., 1969). The uptake of glutamine across the neuronal plasma membrane occurs against the concentration gradient, and is sodium-dependent (e.g. Tamarappoo et al., 1997). The stoichiometry is not known, but uptake of one molecule of glutamine could lead to the entry of 3 Na + (or H+), which would imply the expenditure of one ATP by the Na/K-ATPase. Therefore, one 'transmitter Glu cycle' of vesicular uptake and release, astrocytic uptake and amidation, and neuronal uptake of glutamine, could lead to the use of at least four molecules of ATP per molecule of Glu, two in neurons, and two in astrocytes, in addition to the ATP used for vesicular release. In comparison, complete oxidation of one molecule of glucose to CO2 and water gives 38 molecules of ATE Glutamatergic neurotransmission leads to the consumption of ~ 10% of this energy, since one molecule of glucose is required for the formation of one molecule of Glu. In this calculation we have left out the ATP expenditure inherent in the depolarization of presynaptic membrane which triggers transmitter release and the depolarization of postsynaptic membranes caused by Glu receptor activation. Assuming a flux of transmitter Glu to astrocytes, which is at most 30% of the cerebral TCA cycle rate, we have that <3% of the energy extracted from serum glucose is consumed in the handling of transmitter Glu. The clinical and experimental use of radiolabeled glucose and 2-deoxyglucose in PET studies and autoradiography rests on the assumption that cerebral activity increases the consumption of ATE which in turn necessitates an increase in uptake and metabolism of 10

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glucose. Because we tend to think of cerebral activity as neuronal activity, the view that glucose is primarily taken up by astrocytes implies that a mechanism must exist that couples astrocytic uptake of glucose to neuronal activity. One such coupling mechanism may be the astrocytic uptake and amidation of transmitter Glu, which causes the consumption of two molecules of ATP per molecule of Glu. As shown by Pellerin and Magistretti (1994, 1997), uptake of Glu into cultured astrocytes leads to an increase in Na/K-ATPase activity as well as an increase in glucose uptake and lactate export. This lactate could then be a main metabolic substrate for neurons (reviewed by Magistretti et al., 1999), and it could mediate cerebral vasodilation (Laptook et al., 1988). 2.7. SUMMARY The reviewed literature leaves us with an impression of the cellular interchange of Glu and its precursors and metabolites (Fig.. 2) that is somewhat different from the classical idea of the 'glutamine cycle' with its near-stoichiometric exchange of Glu and glutamine between neurons and astrocytes. (1) Glu is synthesized from glutamine, as previously thought, but also from neuronal precursors supplied by neuronal pyruvate carboxylation. (2) Transmitter Glu is mostly taken up into astrocytes for conversion to glutamine, but an unknown fraction is metabolized by the astrocytic TCA cycle, either fully to CO2 and water, or only partially, to malate which is converted to pyruvate and hence lactate. (3) Uptake of Glu into astrocytes stimulates astrocytic uptake of serum glucose and export of lactate to the extracellular fluid. (4) Lactate, whether formed from transmitter Glu or serum glucose, may increase regional cerebral blood flow and act as a main neuronal energy substrate. (5) Glutamine is shunted to neurons where it to a large extent is metabolized to CO2 and water and, possibly to a lesser extent, is converted to transmitter Glu. (6) The uptake processes related to the handling of transmitter Glu and glutamine together with the formation of glutamine cause the expenditure of ~3% of the total energy of the serum glucose taken up by the brain.

3. ANATOMICAL SYSTEMS 3.1. IS GLUTAMATE IMMUNOLABELING EVIDENCE OF A NEUROTRANSMITTER ROLE FOR GLUTAMATE? The analysis of Glu immunogold-labeled preparations is complicated by the ubiquitous presence of Glu in the CNS. Quantitative analysis of the immunolabeling and comparisons of terminals with other profiles are therefore essential. As biochemical studies have demonstrated high levels of Glu in synaptic vesicles (see above), glutamatergic terminals should be enriched in Glu. Data from immunogold studies support this notion (see Sections 3.2-3.9 and Fig. 6C). However, the ratios of Glu immunolabeling over presumed glutamatergic terminals versus that over other profile types can be rather modest (from less than 2 and higher, depending on what profile types are compared). As the Glu labeling density varies within a population of terminals (due to technical and biological factors such as synaptic vesicle density; Ericson et al., 1995), a proportion of presumed glutamatergic terminal profiles will fall within the range of labeling densities observed over inhibitory terminals (e.g. Broman and Ottersen, 1992; Murphy et al., 1996; Fig. 3). Thus, a relatively low level of Glu immunogold labeling over a certain terminal profile does not preclude Glu as a transmitter in that particular terminal. 11

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'oll Ts

/?-

,

15tl

J

Pr~ I

pl~o"

Fig. 3. (A, B) Electron micrographs from a glutamate immunogold-labeled section of the rat spinal cord dorsal horn showing a presumed primary afferent C fiber terminal (dense sinusoid axon terminal, DSA) in lamina II and a primary afferent terminal (PAT) in lamina III anterogradely labeled (arrow) with choleragenoid-horseradish peroxidase conjugate (B-HRP). Note the higher density of gold particles over the DSA and the PAT in comparison to surrounding tissue and the profile containing pleomorphic vesicles (P) (calibration bar valid for A and B). (C) Histograms demonstrating the distribution of normalized glutamate immunogold labeling densities over different profile types in different laminae of the rat dorsal horn. Vertical lines indicate the average labeling density over laminae III-IV and lamina II, respectively. PATs, B-HRP-labeled primary afferent terminals; DSAs, dense sinusoid axon terminals. (Modified from Broman et al., 1993.)

Is an enrichment of Glu in a population of terminals conclusive evidence of a transmitter role? Many studies have demonstrated enrichment of Glu in nerve terminals defined as glutamatergic on other grounds (see below). However, strictly speaking, the finding of Glu enrichment fulfills only the first of the four main criteria of a neurotransmitter (see Section 1). A critical question is whether Glu may be present in high concentrations in terminals not using Glu as a transmitter. In GABA- and glycine-enriched terminals or terminals displaying 12

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characteristic inhibitory features, low levels of Glu have been detected (e.g. Somogyi et al., 1986; Bramham et al., 1990; Broman et al., 1990, 1993; Todd et al., 1994; 13rnung et al., 1998; however, see Sandler and Smith, 1991; Helfert et al., 1992). The levels of Glu in terminals with other transmitter signatures (e.g. monoaminergic) are largely unknown (but see Torrealba and Mtiller, 1999). However, enrichment of Glu has been detected in cholinergic motor nerve terminals innervating fast-twitch (but not those innervating slow-twitch) muscle fibers (Waerhaug and Ottersen, 1993). It remains to demonstrate if the latter terminals, in addition to acetylcholine, also release Glu. If so, Glu should in this case serve a function other than that of a fast-acting neurotransmitter. In cholinergic terminals in the basal ganglia, Clarke et al. (1997) detected levels of Glu intermediate to those in terminals with asymmetric and symmetric synapses, respectively. A co-release of acetylcholine and Glu has been demonstrated from presumed cholinergic synaptosomes and from cholinergic terminals of the Torpedo electric organ (Docherty et al., 1987; Vyas and Bradford, 1987). Thus, co-localization of Glu with other substances, including classical transmitters, may be compatible with transmitter roles for both substances. However, it cannot be excluded that significant levels of 'metabolic' Glu may be present in certain populations of terminals. In conclusion, although an enrichment of Glu within nerve terminals speaks strongly in favor of a transmitter role for Glu, immunogold data should be interpreted with due caution. 3.2. SPINAL CORD

3.2.1. Primary afferent terminals Since Curtis and Watkins (1960) first observed the excitatory effects of Glu on spinal cord neurons, a long series of biochemical and physiological/pharmacological studies have provided support for a role of Glu as a primary afferent neurotransmitter (reviewed by Rustioni and Weinberg, 1989; Willis and Coggeshall, 1991; Todd and Spike, 1993; Broman, 1994). Important observations include diminished uptake of Glu in the dorsal horn following dorsal rhizotomy and blockage of transmission between primary afferent terminals and dorsal horn neurons with excitatory amino acid receptor antagonists. The literature contains some controversies with respect to the proportion and types of primary afferent terminals that use Glu as a neurotransmitter. Earlier studies emphasized a transmitter role for Glu in large myelinated primary afferent fibers (reviewed by Salt and Hill, 1983). Other findings pointed to a transmitter role for Glu in thinly myelinated and unmyelinated fibers (Schneider and Perl, 1988). De Biasi and Rustioni (1988), using an immunogold technique, were the first to demonstrate the presence of Glu in primary afferent terminals. They detected the presence of Glu in a large proportion of superficial dorsal horn terminals identified by morphological criteria as probable primary afferent terminals. They also noted the presence of Glu in primary afferent terminals traced by anterograde transport or dorsal rhizotomy. Through quantitative evaluation of Glu immunogold labeling, Maxwell et al. (1990a) extended these findings by detecting enrichment of Glu in different types of glomerular central terminals (of probable primary afferent origin) in the superficial dorsal horn. Similar observations were also made by Merighi et al. (1991). Combining anterograde transport of choleragenoid-horseradish peroxidase conjugate (B-HRR injected into dorsal root ganglia) and Glu immunogold labeling, Broman et al. (1993; Fig. 3) demonstrated enrichment of Glu in primary afferent terminals in laminae I and III-V. The levels of Glu in these terminals were 2-3 times higher than the tissue average and 13

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the levels detected in putative inhibitory terminals. A comparison with the labeling density over Glu conjugates in model sections (Ottersen, 1989) processed together with the tissue sections indicated that the B-HRP-labeled terminals contained about 20-25 mM of fixed Glu. Even higher levels of Glu (about 50% higher) were detected in dense sinusoid axon terminals (DSAs) in lamina II. These latter terminals presumably originate from primary afferent C-fibers, whereas those labeled with B-HRP primarily are of myelinated primary afferent origin. The values of Glu labeling density were normally distributed and there was no evidence of a separate population of primary afferent terminals containing low levels of Glu. Valtschanoff et al. (1994) similarly demonstrated enrichment of Glu in B-HRP-labeled primary afferent terminals in the deep dorsal horn. The same authors, using anterograde transport of wheat germ agglutinin-horseradish peroxidase (WGA-HRP), in addition detected high levels of Glu in primary afferent terminals in laminae I-II. Broman and/kdahl (1994) found significant positive correlations between the density of synaptic vesicles and the density of Glu immunogold labeling in DSAs and B-HRP-labeled primary afferent terminals, but not in putative inhibitory terminals. This finding supports a vesicular localization of Glu in primary afferent terminals. The presence of Glu has also been examined in select subpopulations of primary afferent terminals. Maxwell et al. (1993), combining intraaxonal staining with immunogold labeling, detected high levels of Glu in hair follicle afferent boutons. In Clarke's column, giant boutons, likely to originate from primary afferent Ia fibers, are enriched in Glu (Maxwell et al., 1990b), as are Ia afferent terminals (labeled by transganglionic transport of B-HRP) forming synaptic contacts with motoneurons and neurons in the central cervical nucleus ((~rnung et al., 1995). High levels of Glu have also been detected in superficial dorsal-horn primary afferent terminals that were labeled by transganglionic transport of WGA-HRP injected into nipples (Rousselot et al., 1994). In lampreys, high levels of Glu have been detected in primary afferent dorsal column axon terminals (Shupliakov et al., 1992). A strong correlation between Glu immunogold labeling density and the density of synaptic vesicles was evident in these terminals. In conclusion, data from biochemical, physiological/pharmacological and immunocytochemical studies concur in supporting Glu as a primary afferent neurotransmitter. Although it cannot be excluded that a subpopulation of primary afferent terminals that have escaped investigation transmit their signals by other means, the available evidence speaks strongly in favor of Glu as a transmitter in most, if not all, primary afferent terminals (Fig. 4). This of course does not exclude that other compounds, e.g. peptides, are co-released with Glu from selected populations of primary afferent terminals. 3.2.2. Intrinsic n e u r o n s

Glu immunogold data are sparse on terminals that originate from neurons intrinsic to the spinal cord. This reflects the problem of identifying such terminals by morphological criteria or by neuroanatomical tract tracing. There is, however, indisputable physiological evidence of polysynaptic excitatory inputs to neurons in the spinal cord following peripheral stimulation (Willis and Coggeshall, 1991). Application of NMDA and non-NMDA receptor antagonists depresses such responses, indicating a possible role for Glu as a transmitter in synapses of excitatory local-circuit neurons (Broman, 1994). Dorsal horn injections of D-[3H]aspartate labels numerous neurons in the dorsal horn, especially in its superficial part, further supporting the presence of glutamatergic local-circuit neurons in the spinal cord (Rustioni and Cu6nod, 1982; Antal et al., 1991). 14

Biochemistry and anatomy of transmitter glutamate

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Fig. 4. Drawing of the spinal cord depicting pathways and neurons likely to use glutamate as a neurotransmitter. 1 = primary afferent fibers; 2 = local-circuit neurons in the superficial dorsal horn; 3 = corticospinal fibers; 4 -bulbospinal input to the intermediolateral nucleus; 5 = vestibulospinal input to the central cervical nucleus; 6 = the spinocervical tract. For further details, see Section 3.2.

To identify terminals of intrinsic neurons, Todd et al. (1994) used immunolabeling for the peptide neurotensin, which on the basis of lesion studies is believed to have exclusive intraspinal origin. Neurotensin neurons are located primarily around the border between laminae II and III, and a neurotensin-immunoreactive fiber plexus, presumably originating from these neurons, is present in laminae I-II. Todd et al. (1994) demonstrated enrichment of Glu in neurotensin-immunoreactive terminals in laminae I-II, thus providing support for glutamatergic local-circuit neurons in the superficial dorsal horn (Fig. 4). Maxwell et al. (1997) examined the presence of GABA, glycine and Glu in axon terminals of four intracellularly HRP-labeled dorsal-horn interneurons driven by group-II muscle afferents. The terminals of one of these neurons displayed morphological characteristics of inhibitory terminals and were enriched in glycine. The terminals of the other three neurons displayed characteristics of excitatory terminals. Although high levels of Glu were detected in mitochondria in these terminals, cytoplasmic Glu labeling was sparse. The sparse cytoplasmic labeling does not exclude enrichment of Glu in these terminals, as the diaminobenzidine reaction product of the intracellular staining may have suppressed immunoreactivity.

3.2.3. Descending inputs Biochemical studies and retrograde labeling with D-[3H]aspartate provide strong evidence for an excitatory amino acid as transmitter in the corticofugal input to the spinal cord (Storm-Mathisen and Ottersen, 1988; Rustioni and Weinberg, 1989; Fig. 4). That Glu serves as a corticospinal neurotransmitter is further supported by the presence of high levels of Glu in corticospinal terminals in the dorsal horn (Valtschanoff et al., 1993). A large number of descending pathways from the hypothalamus and the brainstem terminate in different regions of the spinal gray matter (Holstege, 1995; Jones, 1995; Tracey, 1995). The effects exerted by these pathways include both excitation and inhibition. Beside the well defined monoaminergic projections from e.g. the raphe nuclei and the locus coeruleus (Broman, 1994), inhibitory amino acids (GABA and glycine) have been detected in terminals of bulbospinal projections to motoneurons (Holstege, 1991; Holstege and Bongers, 1991). 15

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J. B r o m a n et al.

Surprisingly little is known of a possible role of Glu as a transmitter in the descending brainstem pathways. Although negative findings in studies with retrograde transport of D-[3H]aspartate do not exclude a transmitter role of Glu (see Section 1), no positive evidence has emerged from such experiments (Rustioni and Cu6nod, 1982). Several studies have detected the presence of Glu in cell bodies of brainstem neurons projecting to the spinal cord (Beitz and Ecklund, 1988; Mooney et al., 1990; Nicholas et al., 1992; Liu et al., 1995), but cell body labeling for Glu is unreliable as marker for glutamatergic neurons. Positive evidence of a glutamatergic bulbospinal projection has been provided by Ragnarson and collaborators, who demonstrated enrichment of Glu in a proportion of vestibulospinal terminals in the central cervical nucleus (Ragnarson, 1998; Fig. 4). Examination of the spinal cord caudal to spinal transection demonstrates lowered levels of Glu in the sympathetic intermediolateral nucleus, as detected with light microscopical immunocytochemistry (Morrison et al., 1989), and lower than normal density of Glu-immunoreactive terminals on sympathetic preganglionic neurons (Llewellyn-Smith et al., 1997; Fig. 4).

3.2.4. Glutamatergic input to defined spinal neurons The inputs to several different populations of spinal cord neurons (identified by morphological criteria, tract tracing or intracellular staining) have been examined with immunogold labeling for Glu and other compounds. Inherent to the variation of Glu immunogold labeling density within a terminal population (see Section 1), the criteria for defining a terminal as putatively glutamatergic vary somewhat between laboratories. Thus, comparisons of the percentages obtained in different studies should be made with this caution in mind. Murphy et al. (1996) examined terminals that contacted the soma or dendrites of retrogradely labeled motoneurons in the phrenic nucleus. Fifty-five percent of such terminals contained round vesicles and were enriched in Glu. Nerve terminals containing flattened vesicles comprised 42%, and presumably contain inhibitory amino acids. Cell bodies and dendrites displayed similar percentages of round and flattened vesicle terminals. Lindfi and collaborators examined the inputs to large cell bodies of alpha-motoneuron size in the L7 segment (Ornung, 1997). Of the terminals containing spherical vesicles (types S and M), all M terminals and most S terminals were enriched in Glu and displayed a positive correlation between the Glu immunogold labeling density and the density of synaptic vesicles. A proportion of S terminals contained low levels of Glu and displayed no such correlation. C-type (cholinergic) terminals contained levels of Glu intermediate to those detected in terminals with flattened and spherical vesicles, respectively, and displayed no correlation between the densities of synaptic vesicles and gold particles. Between 13.9 and 17.2% of the inputs to neuronal soma and proximal dendrites were classified as glutamatergic. Ornung et al. (1998) examined the inputs to motoneuron dendrites. Thirty-five percent of the boutons were immunopositive for Glu and 59% for GABA and/or glycine. First-order (stem) dendrites differed from more distal dendrites by having a lower proportion of Glu-enriched terminals (18%) and a higher proportion of glycine and/or GABA-immunoreactive terminals (69%). Maxwell et al. (1997) examined inputs to dendrites of group-II spinal interneurons and identified two types of boutons. The first of these bouton types contained spherical vesicles and was enriched in Glu. The second type contained pleomorphic vesicles and was immunoreactive for GABA and/or glycine. Neurons at the origin of several ascending somatosensory pathways have been examined for the presence of glutamatergic inputs. Westlund et al. (1992) investigated inputs to three intracellularly labeled spinothalamic tract neurons in the deep dorsal horn. Of the 16

Biochemistry and anatomy of transmitter glutamate

Ch. I

terminals located on the soma of these neurons, 46.5% were enriched in Glu. The proportion of Glu-immunoreactive terminals on the dendrites of these neurons was slightly higher (50.5%). In the superficial dorsal-horn, Glu-immunoreactive terminals constitute 37% and GABA-immunoreactive terminals 20% of the total terminal length on retrogradely labeled spinothalamic tract cells (Lekan and Carlton, 1995). The percentages were similar on soma and dendrites of these cells. Maxwell et al. (1992, 1995) have examined neurons at the origin of the spinocervical tract and the postsynaptic dorsal column pathway. The percentages of Glu-enriched terminals on these cells were estimated to 42% and 53%, respectively. Llewellyn-Smith et al. (1992, 1998) examined the inputs to sympathetic preganglionic neurons. They detected enrichment of Glu in about two-thirds of the terminals contacting sympathoadrenal neurons and in about half of the terminals contacting neurons which project to the superior cervical ganglion. Most of the remaining terminals were immunoreactive for GABA.

3.2.5. The spinocervical tract The spinocervical tract terminates before reaching the brainstem and will therefore be included in this section. Other somatosensory pathways will be included in the sections below. Spinocervical tract terminals are enriched in Glu, containing about 2.4 times the amount of Glu detected in putative inhibitory terminals in the lateral cervical nucleus (Broman et al., 1990). There is also a significant positive correlation between Glu immunogold labeling density and the density of synaptic vesicles in spinocervical tract terminals, pointing to a vesicular localization of Glu (Kechagias and Broman, 1995). Further, spinocervical tract terminals display high Glu/glutamine ratios, indicating a high rate of Glu synthesis from glutamine (Kechagias and Broman, 1994). There is thus a strong case for Glu as a spinocervical tract neurotransmitter (Fig. 4). 3.3. BRAINSTEM

3.3.1. Medulla oblongata and pons Glu immunogold labeling have been used to examine different types of primary afferent terminals in different loci of the brainstem. In the cuneate nucleus, terminals of large-caliber somatic sensory primary afferent fibers are enriched in Glu (De B iasi et al., 1994a), corroborating the observations on such terminals in the deep dorsal horn of the spinal cord (Broman et al., 1993; Valtschanoff et al., 1994). Saha et al. (1995a) examined vagal afferent terminals in the solitary tract nucleus and the dorsal vagal motor nucleus, and detected enrichment of Glu in 57% of such terminals. However, they considered terminals to be immunoreactive for Glu only if their labeling density exceeded the mean tissue labeling density with 2.576 times the standard deviation. Indeed, Sykes et al. (1997) examined the same population of terminals and detected higher-than-average tissue level of Glu in all examined vagal afferent terminals. Hackney et al. (1996) examined type-I cochlear afferent terminals and detected enrichment of Glu, high Glu/glutamine ratios and partly Ca2+-dependent depletion of Glu following K+-induced depolarization in such terminals. In lamina II of the caudal spinal trigeminal nucleus, glomerular terminals of presumed primary afferent origin are enriched in Glu (Iliakis et al., 1996). Thus, similar to the situation in the spinal cord (Section 3.2.1), enrichment of Glu is evident in different types of primary afferent terminals in different loci of the brainstem (Fig. 5). 17

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J. Broman et al.

7

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Fig. 5. Drawing of the brainstem depicting neurons and pathways likely to use glutamate as a neurotransmitter. 1 -- primary afferent inputs to the dorsal column nuclei (a), the solitary tract nucleus (b), and the cochlear nucleus (c); 2 = granule cell/parallel fibers in the dorsal cochlear nucleus" 3 --- calyces of Held in the medial nucleus of the trapezoid body" 4 -- cochlear nucleus inputs to the lateral superior olive; 5 = input to the oculomotor nucleus from the ventral lateral vestibular nucleus; 6 = input to the oculomotor nucleus from the abducens nucleus; 7 -corticocollicular inputs; 8 = spinal input to the periaqueductal gray; 9 = inputs to the red nucleus and pontine nuclei from the cerebellar nuclei. For further details, see Section 3.3.

In the dorsal column nuclei, in addition to primary afferent terminals also terminals of cortical origin are enriched in Glu (Rustioni and Weinberg, 1989; Valtschanoff et al., 1991), corroborating observations of retrograde transport of o-[3H]aspartate in corticocuneate fibers (Rustioni and Cu6nod, 1982). However, only low levels of Glu have been detected in most terminals of dorsal horn neurons projecting to the dorsal column nuclei (the postsynaptic dorsal column pathway, PSDC; De Biasi et al., 1995). This finding suggests that the PSDC, in contrast to other ascending somatosensory pathways (Sections 3.2.5, 3.3.2 and 3.5), do not use Glu as a neurotransmitter. Enrichment of Glu has also been detected in cortical afferent terminals in the solitary tract nucleus (Torrealba and Mtiller, 1996, 1999). The latter and a previous study (Saha et al., 1995b) estimated similar proportions of GABA-immunoreactive terminals in the solitary tract nucleus (36 and 33%, respectively), but differ in their estimates of the proportion of putative glutamatergic terminals (61 and 40%, respectively). Neurons in the solitary tract nucleus are retrogradely labeled following injections of D-[3H]aspartate into the ventrolateral medulla, thus supporting a role for excitatory amino acids as neurotransmitters in this pathway (Somogyi et al., 1989). In the area postrema, Walberg and Ottersen (1992) detected a large number of GABA-immunoreactive terminals and high levels of Glu in a substantial proportion of terminals. High proportions of GABA-immunoreactive and Glu-enriched terminals have also been observed in the trigeminal motor nucleus (Yang et al., 1997). In addition to cochlear primary afferents (see above), other nerve terminal populations in the auditory system have been subjects for analysis with Glu immunolabeling. Grandes and Streit (1989) examined calyces of Held (originating from the contralateral ventral cochlear 18

Biochemistry and anatomy of transmitter glutamate

Ch. I

nucleus) in the medial nucleus of the trapezoid body and detected high levels of Glu and accumulation of gold particles over clusters of vesicles and mitochondria in such terminals (Fig. 5). Helfert et al. (1992) detected high levels of Glu in terminals containing round synaptic vesicles in the lateral superior olive. These terminals presumably originate from the ipsilateral cochlear nucleus (Fig. 5). However, they also detected high levels of Glu in glycine-enriched terminals containing flattened vesicles (though somewhat lower levels of Glu than in round vesicle terminals), presumably originating from the contralateral medial nucleus of the trapezoid body. The functional significance of this co-localization of glycine and Glu is unclear. In the dorsal cochlear nucleus, parallel fiber terminals originating from granule cells are enriched in Glu which is depleted by depolarization with high [K+] in slice experiments (Osen et al., 1995; Fig. 5). High levels of Glu in auditory nerve terminals and granule cell terminals were also reported by Rubio and Juiz (1998), who in addition detected high Glu levels in large 'mossy' terminals in the dorsal cochlear nucleus. There is strong biochemical support for Glu as a neurotransmitter in the massive cortical input to the pontine nuclei (Storm-Mathisen and Ottersen, 1988; Ottersen, 1991). Glu immunogold studies further support this by demonstrating, in the pontine nuclei, enrichment of Glu in terminals with round vesicles (Border and Mihailoff, 1991; Aas et al., 1992) and in terminals labeled by tracer injections into the posterior cingulate cortex (Azkue et al., 1995). In the pontine nuclei and in the nucleus reticularis tegementi pontis, enrichment of Glu has also been demonstrated in terminals originating from the cerebellar nuclei (Schwarz and Schmitz, 1997; Fig. 5). K+-induced depolarization of tissue slices results in CaZ+-dependent depletion of Glu in round vesicle-containing terminals forming asymmetric synaptic contacts in the pontine nuclei (Aas et al., 1992). 3.3.2. Midbrain

Neurons in different regions of the cerebral cortex, including visual cortices, are retrogradely labeled by injections of D-[3H]aspartate into the superior colliculus (Matute and Streit, 1985). Further evidence for Glu as corticocollicular transmitter comes from biochemical studies (Storm-Mathisen and Ottersen, 1988) and the demonstration of enrichment of Glu in corticocollicular terminals (Ortega et al., 1995; Mize and Butler, 1996; Fig. 5). Also retinotectal fibers are labeled by D-[3H]aspartate injections into the superior colliculus (Matute and Streit, 1985) and their terminals are enriched with Glu (Ortega et al., 1995; Mize and Butler, 1996). Enrichment of Glu has also been detected in retinal terminals in the pretectum (Nunes-Cardoso et al., 1991) and in the optic tectum of Vipera (Reperant et al., 1997). There is thus strong support for Glu as a neurotra'nsmitter in the visual inputs to the midbrain. That also somatosensory inputs to the midbrain may use Glu as a neurotransmitter is supported by the detected enrichment of Glu in the spinal input to the periaqueductal grey (Azkue et al., 1998; Fig. 5). As the input from the lateral cervical nucleus to the midbrain essentially are collaterals of cervicothalamic tract fibers, the demonstration of Glu enrichment in cervicothalamic tract terminals (Broman and Ottersen, 1992; Kechagias and Broman, 1995) provides indirect evidence for Glu as a cervicomesencephalic neurotransmitter. Similar to other projections from the cerebellar nuclei (except those to the inferior olive which are GABAergic), terminals of cerebellar origin in the red nucleus are enriched in Glu (Schwarz and Schmitz, 1997; Fig. 5). Enrichment of Glu has also been detected in terminals in the oculomotor nucleus originating from the abducens and ventral lateral vestibular nuclei (Nguyen and Spencer, 1999; Fig. 5). 19

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3.4. C E R E B E L L U M An extensive review of the chemoarchitecture and anatomy of the cerebellum was published in a recent volume of the Handbook Series (Voogd et al., 1996). This volume should be consulted for a complete bibliography. Another recent review is that of Ottersen and Walberg

(2ooo). The major afferent pathways to the cerebellar cortex are the mossy and climbing fiber systems. Climbing fibers establish direct contact with the Purkinje cells, whereas the mossy fibers influence the Purkinje cells indirectly, through the granule cell-parallel fiber system. The excitatory nature of each of these systems (including the parallel fibers) is well-established (Ito, 1984) and several lines of evidence point to Glu as their likely transmitter (Fig. 6). As for the mossy fibers, these have been shown to display a strong Glu immunogold signal (Somogyi et al., 1986) whose intensity is positively correlated to the packing density of synaptic vesicles (Ji et al., 1991). The Glu immunolabeling of these terminals was abolished by depolarization of cerebellar slices with high [K +] (Ottersen et al., 1990). This implies that the immunolabeling is likely to represent a transmitter pool of Glu, rather than a metabolic pool unrelated to synaptic transmission. The postsynaptic elements of the mossy fibers express several types of glutamate receptor, including N M D A and AMPA receptors (Cox et al., 1990; Gallo et al., 1992; Petralia and Wenthold, 1992), and pharmacological data are consistent with the idea that glutamate acts as their endogenous ligand (Garthwaite and Brodbelt, 1990). The mossy fibers also show strong immunoreactivity for phosphate-activated glutaminase, the major glutamate-synthesizing enzyme (Fig. 6E). The evidence referred to above must not be taken to exclude the possibility that some mossy fibers use other signal substances, instead of or in addition to Glu. Notably, subpopulations of mossy fibers contain choline acetyl transferase (CHAT; Kan et al., 1978), the enzyme responsible for acetylcholine synthesis, although there is still conflicting evidence as to the extent of the cholinergic input to the cerebellum (Ottersen and Walberg, 2000). Mossy fibers also express several neuroactive peptides that may serve modulatory functions (Voogd et al., 1996).

Fig. 7. Immunocytochemical analyses show distinct cellular and subcellular compartmentation of Glu, glutamine, and the enzyme (phosphate-activated glutaminase; PAG) responsible for the conversion of precursor glutamine to Glu. (A) Light microscopical distribution of glutamine in the cerebellar cortex. (Modified from Nagelhus et al., 1996.) Strong glutamine-like immunoreactivity is found in the cell bodies (double arrowheads) and radial processes (arrowheads) of the Golgi epithelial cells (a class of astrocyte). Labeled astrocytes (crossed arrows) also occur in the granule cell layer (gr). Purkinje cells (arrows) are weakly stained; mo, molecular layer. The animal was subjected to hypo-osmotic stress to increase the concentration of glutamine in glial cells (Nagelhus et al., 1996) (scale bar 25 Ixm). (B) Electron micrograph of ultrathin section double labeled for glutamine (large gold particles) and Glu (small particles). The ratio between the two particle sizes differs significantly between glial cells (g; an astrocyte in the granule cell layer) and granule cells (gr; a type of putative glutamatergic cell) (scale bar 0.4 Ixm). (C) Double-labeled section similar to that in B. Large particles signaling glutamine predominate in astrocytes (upper right) while small particles (signaling Glu) are concentrated in mossy fiber terminals (mf) and particularly in the mitochondria of these. Relatively high densities of small particles also occur in the dendritic digits (d) of the granule cells while terminals of the putative GABAergic Golgi cells (asterisk) are weakly labeled (scale bar 0.4 Ixm). (D) A control section that had accompanied the tissue sections through the double-labeling procedure shows selective accumulation of large and small particles over glutamine conjugates and Glu conjugates, respectively. GABA conjugates or brain macromolecules treated with glutaraldehyde in the absence of any free amino acid ('none') were almost devoid of immunolabeling (scale bar 0.5 Ixm). (E) Electron micrograph of ultrathin section incubated with an antibody to PAG (Laake et al., 1999). A mossy fiber terminal (mf) contacting a unipolar brush cell (UBC) displays strong immunolabeling confined to mitochondria (cf. C) (scale bar 1 I~m). 20

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The climbing fibers are now believed to use glutamate as transmitter, like the major proportion of mossy fibers. They are strongly enriched in glutamate (Ottersen et al., 1992) and their terminals are apposed to Purkinje cell thorns that exhibit a high density of AMPA receptors (Landsend et al., 1997). L-aspartate was long considered the most probable transmitter candidate in climbing fibers (for review see Ottersen and Walberg, 2000). Experimental support of this view was provided by Toggenburger et al. (1983) and Vollenweider et al. (1990), who showed that the evoked release of endogenous aspartate from slices of the cerebellar cortex could be reduced by lesions of the inferior olive by 3-acetylpyridine. However, quantitative immunogold analysis of perfusion-fixed brains (Zhang et al., 1990) revealed only a low level of L-aspartate immunoreactivity in the climbing fiber terminals. Notably, the nerve cell bodies in the inferior olive, which give rise to the climbing fibers, displayed a significantly stronger L-aspartate immunogold signal than the latter. These observations were difficult to reconcile with a transmitter role of L-aspartate. False-positive results in slice experiments such as those above could ensue due to temporary energy failure during the preparation of the slice, which would cause a buildup of L-aspartate in nerve terminals that contain only sparse amounts of this amino acid under physiological conditions (Gundersen et al., 1998). On the other hand, false-negative results in immunogold analyses would occur if the L-aspartate pool were released during the initial stage of the perfusion (prior to irreversible fixation by glutaraldehyde) or if transmitter L-aspartate were inaccessible to immunogold detection. The latter explanations are unlikely but cannot be excluded. The observation that climbing fibers are able to take up and transport D-[3H]aspartate (Wiklund et al., 1984) does not help elucidate this problem since glutamate transporters do not differentiate between D-aspartate (used as exogenous tracer) and the endogenous amino acids L-aspartate and L-glutamate (Chapter 8, this volume). Lesions of the inferior olive have been shown to reduce the evoked release of homocysteic acid from cerebellar slices (Vollenweider et al., 1990). Homocysteic acid is a sulfur-containing excitatory amino acid that has been proposed as a transmitter candidate in several fiber systems, including the climbing fibers (Cu6nod et al., 1989). However, with the advent of specific antibodies it could be demonstrated that homocysteic acid was restricted to glial elements of the cerebellar cortex (Grandes et al., 1991). This was confirmed in independent immunogold experiments (Zhang and Ottersen, 1992). Homocysteic acid is therefore an unlikely transmitter candidate in climbing fibers, although the possibility remains that it is released from glia as part of a complex signaling process (Do et al., 1997). If glutamate or another substrate of glutamate transporters is responsible for climbing fiber neurotransmission one would expect an interference with glutamate transport to affect the postsynaptic response to climbing fiber activation. Following this line of reasoning, Takahashi et al. (1996) injected D-aspartate into Purkinje cells, in an attempt to inhibit postsynaptic glutamate uptake at sites of afferent excitatory input. The D-aspartate injections were found to prolong the excitatory postsynaptic current at climbing fiber synapses. This is consistent with the idea that the transmitter is a substrate of Purkinje cell glutamate transporters and that these normally contribute to transmitter removal. Two glutamate transporters (EAAT3 and EAAT4) have been localized to Purkinje cells and one of these (EAAT4) shows a highly specific compartmentation in the postsynaptic membrane (Chapter 8, this volume). The parallel fibers ~ the axons of granule cells - - constitute the third major excitatory fiber system in the cerebellum. The parallel fibers establish synapses with Purkinje cell spines as well as with dendritic stems of interneurons (Palay and Chan-Palay, 1974). Several lines of evidence point to glutamate as the most likely transmitter in the parallel fiber system. 22

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Young et al. (1974) made the seminal observation that a granule cell loss (induced by virus infection) caused a reduction in glutamate content and glutamate/aspartate uptake in the cerebellar cortex. This pioneering study was followed by a series of reports from different laboratories that concluded that content, uptake, as well as release of glutamate depend on intact granule cells (for reviews see: Ito, 1984; Ottersen and Storm-Mathisen, 1984a). With the introduction of the post-embedding immunogold technique it was shown that the parallel fibers were enriched in glutamate (Somogyi et al., 1986; Ottersen, 1987) and that their glutamate content could be depleted by high [K +] as expected of a transmitter pool (Ottersen et al., 1990). Immunogold studies have also established that the postsynaptic specializations of parallel fiber synapses are equipped with AMPA and 32 glutamate receptors (Baude et al., 1994; Nusser et al., 1994; Landsend et al., 1997). The latter receptor, whose biological role is still unclear, is restricted to parallel fiber synapses with Purkinje cell spines. Parallel fiber synapses on dendritic stems appear to be devoid of this receptor (Landsend et al., 1997). The available neurochemical and immunocytochemical data make a very strong case for glutamate as the parallel fiber transmitter. However, some uncertainty remains as to how the transmitter is replenished. Compared to the mossy fibers (see above), the parallel fibers display rather low immunoreactivity for PAG (Laake et al., 1999) and are thus likely to depend on alternative sources for transmitter replenishment (see Section 2). 3.5. THALAMUS An extensive review of the anatomy, physiology and neurochemistry of the thalamus has recently been published in this Handbook Series (Jones, 1998). The present review focuses on the main excitatory inputs to the thalamus, i.e. the principal subcortical afferents and the corticothalamic input.

3.5.1. Corticothalamic projections The most well established glutamatergic projection to the thalamus is the massive corticothalamic input originating from neurons in lamina VI of the cerebral cortex (Fig. 7). The glutamatergic nature of this input is supported by biochemical findings, retrograde transport of D-[3H]aspartate as well as by physiological/pharmacological data (Storm-Mathisen and Ottersen, 1988; Rustioni and Weinberg, 1989; Ottersen, 1991; Broman, 1994). Immunogold studies of the thalamus support this conclusion by demonstrating enrichment of Glu in terminals with morphologic features of corticothalamic terminals (Montero and Wenthold, 1989; Montero, 1990; Hamori et al., 1990; Broman and Ottersen, 1992; De Biasi et al., 1994b; Ericson et al., 1995; Blomqvist et al., 1996).

3.5.2. Principal subcortical afferents During the last decade increasing evidence has been gathered for a transmitter role of Glu also in the principal subcortical afferents to the thalamus. Although thalamic injections of D-[3H]aspartate do not label neurons of ascending somatosensory pathways (Rustioni et al., 1983), pharmacological data strongly suggested a role for excitatory amino acid receptors in the synapses between sensory afferents and thalamic neurons (reviewed by Salt and Eaton, 1996). The findings in Glu immunogold studies support this view by universally demonstrating enrichment of Glu in sensory afferent terminals. Montero and Wenthold (1989) and Montero 23

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Fig. 7. Schematic drawing of a transverse section through the forebrain depicting pathways likely to use glutamate

as a neurotransmitter. 1 -- principal subcortical afferents to the thalamus from somatosensory relay nuclei and the spinal cord (a), cerebellar nuclei (b), and retina (c); 2 --- intrinsic neurons and retinal inputs to the hypothalamus; 3 = thalamocortical inputs; 4 = corticothalamic inputs; 5 = cortical inputs to the basal ganglia and other areas in the brainstem and spinal cord; 6 = associational and commissural connections in the cerebral cortex. For further details, see Sections 3.5-3.9.

(1990) detected high levels of Glu in morphologically identified retinal terminals in the lateral geniculate nucleus of monkeys and cats. In the ventrobasal complex, enrichment of Glu has been detected in sensory afferent terminals identified either by morphological criteria (Hamori et al., 1990) or by anterograde transport (cervicothalamic tract: Broman and Ottersen, 1992; afferents from the dorsal column nuclei: De Biasi et al., 1994a). Spinothalamic tract terminals in the cat nucleus submedius (Ericson et al., 1995) and in the posterior thalamic region of monkeys (Blomqvist et al., 1996) are similarly enriched in Glu. Enrichment of Glu has also been detected in cerebellothalamic terminals in the ventromedial and ventrolateral nuclei (Schwarz and Schmitz, 1997). In all studies in which both sensory afferent terminals and terminals of presumed cortical origin have been examined, the levels of Glu are higher in the corticothalamic than in the sensory afferent terminals. The higher density of synaptic vesicles in the corticothalamic terminals may explain this. In support of a vesicular localization of Glu in sensory afferent terminals, positive correlation between the Glu immunogold labeling density and the density of synaptic vesicles has been detected in retinothalamic (Montero and Wenthold, 1989), cervicothalamic (Kechagias and Broman, 1995) and spinothalamic tract terminals (Ericson et al., 1995; Blomqvist et al., 1996). In conclusion, data from immunogold studies concur with pharmacological findings in support of Glu as a transmitter in the principal subcortical afferents to the thalamus (Fig. 7). 3.6. HYPOTHALAMUS Earlier investigations focused on peptides and other slow-acting agents as the principal neuroactive substances in neuronal circuits of the hypothalamus (Swanson, 1987). However, Van den Pol (1991) examined the presence Glu in several medial hypothalamic nuclei (the suprachiasmatic, arcuate, ventromedial, supraoptic and parvocellular and magnocellular periventricular nuclei) and detected a population of boutons displaying strong immunoreactivity for Glu. Decavel and Van den Pol (1992) similarly found Glu-enriched terminals forming synapses with hypothalamic neurosecretory neurons (labeled by intravenous injections of 24

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horseradish peroxidase) in the paraventricular and arcuate nuclei. Quantitative analysis indicates that Glu-enriched terminals constitute 20-40% of all terminals in the supraoptic nucleus and areas surrounding it (Meeker et al., 1993; E1 Majdoubi et al., 1996). In the neural lobe of the pituitary, clusters of small clear vesicles in neurosecretory endings (originating from vasopressin/oxytocin neurons in the hypothalamus) are strongly labeled for Glu (Meeker et al., 1991). These data, together with findings in functional studies (Van den Pol, 1991; Van den Pol and Trombley, 1993), point to Glu as a central mediator of excitation in the hypothalamus. The origin of hypothalamic glutamatergic terminals is presumably diverse, including intrahypothalamic connections (Van den Pol and Trombley, 1993) as well as projections from outside the hypothalamus (Fig. 7). Consistent with the observations on retinothalamic terminals (Section 3.5), enrichment of Glu has been detected in terminals of retinal origin in different hypothalamic nuclei and in different species (Castel et al., 1993; De Vries et al., 1993; Chen and Pourcho, 1995). Hypothalamic glutamatergic systems are modified during changes in neurohormonal status. E1 Majdoubi et al. (1997) found that the supraoptic nucleus increased 40% in size in lactating compared to virgin rats without a concomitant decrease in the volume densities of synapses of Glu-enriched or GABA-immunoreactive terminals, thus demonstrating increases in the total numbers of such synapses. At least part of this increase in the number of glutamatergic and GABAergic synapses in lactating rats is explained by a larger incidence of terminals forming more than one synapse. 3.7. BASAL GANGLIA The basal ganglia, comprising the caudate nucleus, the putamen, the nucleus accumbens, the globus pallidus, the subthalamic nucleus and the substantia nigra, have profuse and complex fiber connections with each other as well as with several other regions of the central nervous system (for some reviews see: Parent, 1990; Parent and Hazrati, 1993, 1995a,b; Gerfen and Wilson, 1996; Levy et al., 1997). It is well established that the basal ganglia play a crucial role in the pathology of many neurological diseases, such as, for instance, Parkinson's disease and Huntington's chorea. In spite of this, in only a minority of the many fiber connections of the basal ganglia has the transmitter substance been identified by means of combined tracing and immunocytochemical studies at the ultrastructural level. Admittedly, a correlation of information gained from various types of investigations has led to the identification of a probable transmitter substance in several of the fiber connections of the basal ganglia. The massive corticostriatal projection emphasizes this point. Early electrophysiological (Kitai et al., 1976; Wilson, 1986) and neurochemical (Spencer, 1976; Divac et al., 1977; Kim et al., 1977; Streit, 1980; Fonnum et al., 1981) investigations were suggestive of a glutamatergic excitatory input to the striatum from the cerebral cortex. A subsequent light microscopical immunohistochemical study documented that a very large number of fibers and bouton-like structures in striatum of the rat display Glu-like immunoreactivity (Glu-LI) (Ottersen and Storm-Mathisen, 1984b). In a more recent electron microscopical study in the rat it was shown that striatal boutons with the ultrastructural characteristics of cortical afferents were enriched in Glu and that they also had a high-affinity uptake of aspartate (Gundersen et al., 1996). Thus, although a combined immunocytochemical investigation of anterogradely labeled corticostriatal boutons has yet to be published, the overall data combined, including the many reports on the differentiated distribution of various types of Glu receptors (see Petralia et al., this volume) sustain the notion that corticostriatal fibers use Glu as a transmitter substance (Fig. 7). It should be noted, however, that the 25

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organization of this projection is very complex, and it still remains an open question whether all corticostriatal fibers, or only a certain percentage of them, use Glu as a transmitter. In a recent light microscopical investigation it was shown that 52-61% of retrogradely labeled corticostriatal neurons displayed Glu-LI (Bellomo et al., 1998). Up to 96% of these neurons were immunopositive when antisera against Glu and aspartate were used simultaneously, and the Glu- and Asp-immunopositive cortical neurons appeared to be largely segregated (Bellomo et al., 1998). The cerebral cortex also sends fibers to other basal ganglia than the striatum, although on a much smaller scale. Thus, the subthalamic nucleus, STN, receives fibers from wide cortical areas (Kunzle and Akert, 1977; Kunzle, 1978; Rinvik et al., 1979; Afsharpour, 1985; Canteras et al., 1988), and the axon terminals have ultrastructural characteristics of other cortical efferent projections (Romansky et al., 1979). Electrophysiological investigations had already provided strong evidence for an excitatory input to the STN from the cerebral cortex (Kitai and Deniau, 1981; Rouzaire-Dubois and Scarnati, 1987; F6ger and Mouroux, 1991; Fujimoto and Kita, 1993), and in a combined tracing and immunocytochemical study in the rat it was indeed shown that a considerable number of the corticosubthalamic boutons are enriched in Glu (Bevan et al., 1995). On a far smaller scale than the prominent corticosubthalamic projection, anterogradely labeled corticopallidal terminals have been identified in the rat (Naito and Kita, 1994a). Although no immunocytochemical correlation was made, these corticopallidal boutons had an ultrastructural appearance similar to cortical efferents in other basal ganglia, and it appears highly likely that these afferents also are glutamatergic. Whether corticopallidal projections exist in other species remains to be settled, but boutons establishing synapses similar to the corticopallidal ones in the rat have been described in the cat (Okoyama et al., 1987) and the monkey (DiFiglia et al., 1982). Whether the substantia nigra receives a direct cortical input is still an open question, although a sparse projection has been demonstrated in the rat (Naito and Kita, 1994b). The thalamus represents the second largest source of afferents to the basal ganglia (for references see De las Heras et al., 1997). In a combined tracing and immunocytochemical study in the rat, it was shown that axon terminals in the subthalamic nucleus from the parafascicular nucleus of the thalamus are highly enriched in Glu (Bevan et al., 1995), lending support to earlier physiological and pharmacological investigations (Mouroux and F6ger, 1993; F6ger et al., 1997). As far as the massive thalamostriatal projection is concerned, the identity of the transmitter substance remains to be determined with certainty (De las Heras et al., 1997). Based on the ultrastructural appearance of the thalamostriatal synapses (Sadikot et al., 1992), however, and combined with the many electrophysiological studies which have shown that the thalamus exerts an excitatory effect on striatal neurons (Purpura and Malliani, 1967; Buchwald et al., 1973; Kitai et al., 1976), it appears highly likely that at least part of the massive and complex thalamostriatal projection is glutamatergic. In a recent light microscopical study it was suggested that many afferents to the ventral striatum from the amygdala in the rat might be glutamatergtic (McDonald, 1996). In the last decade the subthalamic nucleus has taken a central stage in attempts to explain the pathophysiology of Parkinson's disease (Albin et al., 1989b). Although several observations clearly indicate that the original model was too simplified (Marsden and Obeso, 1994; Levy et al., 1997; Obeso et al., 1997), it remains unquestionable that hyperactivity of the subthalamic neurons is a prominent feature in Parkinson's disease. The main efferent projections of the subthalamic nucleus are to both segments of the globus pallidus in primate, the globus pallidus and entopeduncular nucleus in rodents, both the pars compacta 26

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and pars reticulata of the substantia nigra, and a smaller contingent to the tegmental pedunculopontine nucleus (PPN) (for references see the review by Parent and Hazrati, 1995b). Light microscopical immunohistological studies have shown that practically all cells in the STN display strong Glu-LI in the rat (Ottersen and Storm-Mathisen, 1984a,b), in the monkey (Smith and Parent, 1988) and the cat (Albin et al., 1989a) Since the presence of Glu in neuronal cell bodies does not necessarily imply that Glu is used as transmitter, these findings are not decisive, although they accord with electrophysiological data (Kitai and Kita, 1987). More specific support in favor of a transmitter role of Glu was obtained in combined tracing and immunocytochemical studies in the cat which showed that boutons of subthalamonigral fibers are enriched in Glu (Rinvik and Ottersen, 1993). Similar investigations have not been undertaken for the subthalamopallidal or subthalamoentopeduncular projections. On the other hand, in the rat there is ample evidence that the subthalamofugal fibers send branches to both the substantia nigra and the globus pallidus (Van der Kooy and Hattori, 1980). In the primate, neurons of the subthalamic nucleus are segregated in subpopulations according to their target structure (Parent and Smith, 1987; Parent and Hazrati, 1995b). In an immunocytochemical investigation in the monkey it was shown that a certain percentage of boutons that establish asymmetrical synapses with somata and dendrites in the external as well as the internal part of the pallidum display Glu-LI, and that they, furthermore, have a differential distribution in the two pallidal segments (Shink and Smith, 1995). When correlated with the above-mentioned immunohistochemical and tracing studies, as well as with electrophysiological investigations (Robledo and F6ger, 1990), it appears highly likely that the subthalamic projection to both segments of the globus pallidus (including the rodent's entopeduncular nucleus) are glutamatergic. The nature of the transmitter substance in the projection from the subthalamic nucleus to the pedunculopontine nucleus (PPN) remains to be determined. On the other hand, recent investigations have provided a more detailed knowledge of the efferent projections of the PPN. This heterogeneous structure projects profusely upon the basal ganglia, and particularly the subthalamic nucleus and the pars compacta of the substantia nigra, and to a minor degree upon the pallidum and the striatum (Lavoie and Parent, 1994b). A considerable number of cells in the PPN display Glu-LI (Lavoie and Parent, 1994a) and a light microscopical study in the monkey has shown that a portion - - but not all m of the cells projecting to the substantia nigra from the PPN display Glu-LI (Lavoie and Parent, 1994c). In combined tracing and immunocytochemical studies in the rat it was shown that the PPN, or at least a part of it, sends Glu-enriched fibers to the STN (Bevan and Bolam, 1995), and to the entopeduncular nucleus (Clarke et al., 1997). The latter authors could, furthermore, demonstrate that a significant portion of labeled axon terminals from the PPN displayed high levels of immunoreactivity against both Glu and choline acetyltransferase, suggestive of a co-localization of Glu and acetylcholine. 3.8. RETINA The retina has a highly ordered and layered anatomical arrangement and contains five major classes of neurons: photoreceptors (rods and cones), bipolar cells, horizontal cells, amacrine cells and ganglion cells. Bipolar cells form the 'through' pathway between photoreceptors and ganglion cells, the output cells of the retina, whereas the horizontal and amacrine cells form lateral connections. There is strong support for Glu as a neurotransmitter in the 'through' pathway (for references to earlier literature, see Ehinger and Dowling, 1987; Massey and Redburn, 1987; Daw et al., 1989). Glu is released from photoreceptors (Copenhagen and 27

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Jahr, 1989), and several immunocytochemical studies have reported high levels of Glu in photoreceptor cells (Davanger et al., 1991; Kalloniatis and Fletcher, 1993; Jojich and Pourcho, 1996; Huster et al., 1998). Detailed analyses of Glu immunogold labeling of the human and baboon retinae also demonstrate higher concentrations of Glu in the photoreceptor terminals than in the outer parts of both rods and cones (Davanger et al., 1994a). Also bipolar cells have universally been found to contain high levels of Glu (Ehinger et al., 1988; Davanger et al., 1991; Martin and Grfinert, 1992; Kalloniatis and Fletcher, 1993; Jojich and Pourcho, 1996). As for photoreceptor cells, higher concentrations of Glu are evident in bipolar cell terminals than in their cell bodies (Davanger et al., 1994b). Ganglion cells project to several loci of the brainstem and thalamus, and their terminals are enriched in Glu (see Sections 3.3.2, 3.5 and 3.6). Immunocytochemical evidence thus concurs with other data in support of Glu as a transmitter in all synapses of the 'through' pathway from photoreceptors to neurons in the brainstem and thalamus. In the human retina, Glu is co-localized with homocysteic acid in photoreceptor terminals and with glycine in a large subpopulation of bipolar cell terminals (Davanger et al., 1994a,b). In the retina, Glu functions as an inhibitory transmitter in certain photoreceptor-bipolar cell synapses. When light hyperpolarizes the photoreceptor, the release of Glu in the photoreceptor-bipolar cell synapse diminishes. This results in inhibition of off-center bipolar cells, as expected from diminished release of an excitatory transmitter, whereas on-center bipolar cells are excited (Copenhagen, 1991). The Glu excitation of off-center bipolar cells is due to activation of ionotropic Glu receptors, whereas the Glu-induced inhibition of on-center bipolar cells is dependent on metabotropic (mGluR6) receptors (Nakajima et al., 1993; Euler et al., 1996; Sasaki and Kaneko, 1996; Brandstatter et al., 1997; Vardi and Morigiwa, 1997; DeVries and Schwartz, 1999; Morigiwa and Vardi, 1999). 3.9. CEREBRAL CORTEX The role of Glu as a neurotransmitter of neurons in the cerebral cortex has been extensively investigated (Storm-Mathisen and Ottersen, 1988; Tsumoto, 1990; Ottersen, 1991; McCormick, 1992) and a comprehensive discussion of this issue is beyond the scope of the present review (the hippocampus will not be dealt with here, for recent references see Gundersen et al., 1998). As indicated in the previous sections, there is strong support from biochemical, pharmacological and immunocytochemical studies that layers V and VI pyramidal cells which project to subcortical structures use Glu as a neurotransmitter (Fig. 7). Retrograde tracing with D-[3H]aspartate also supports a role for Glu as a neurotransmitter in pyramidal neurons projecting to other areas of the ipsilateral or contralateral cortex (associational and commissural connections; Streit, 1980; Barbaresi et al., 1987; Elberger, 1989; Fig. 7). Pyramidal neurons in the different cortical layers emit local axon collaterals in addition to their longer projections (Somogyi et al., 1998). Small injections of D-[3H]aspartate into different layers of the cortex consequently label pyramidal neurons with vertical or horizontal axon collaterals to the injected area (Kisvarday et al., 1989; Johnson and Burkhalter, 1992). The cerebral cortex also contains local-circuit neurons of different types and in different layers, the majority of which are inhibitory and immunoreactive for GABA (Somogyi et al., 1998). However, a special type of interneuron in layer IV, the spiny stellate neuron, is excitatory and assumed to use Glu as a neurotransmitter (Saint Marie and Peters, 1985; Tsumoto, 1990; Anderson et al., 1994). Thus, there is convincing evidence that excitatory amino acids, most likely Glu, act as neurotransmitter in most projection neurons of the cerebral cortex and presumably also in the spiny stellate local-circuit neurons. 28

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Although Glu is believed to serve a neurotransmitter role in the major thalamic inputs to the cerebral cortex, the evidence in favor of such a role has been less substantial than for the cortical pyramidal cells. The organization of thalamocortical inputs is highly complicated and can be subdivided in different ways (reviewed by Castro-Alamancos and Connors, 1997). The specific projections arise in the primary sensory or motor relay nuclei (e.g. the ventroposterior nucleus, the ventrolateral nucleus and the dorsal lateral geniculate nucleus) and terminate primarily in the middle layers of their corresponding cortical areas. Projections from many other nuclei, e.g. the intralaminar nuclei, the anterior nuclei and the ventromedial nucleus, are usually more diffuse and terminate in either the superficial or deep cortical layers. The latter thalamocortical projections have been referred to as non-specific or unspecific. Injections of D-[3H]aspartate into the cerebral cortex have yielded different results with respect to retrograde labeling of thalamic neurons. In some studies no or only few retrogradely labeled neurons have been detected in the thalamus (Streit, 1980; Baughman and Gilbert, 1981; Barbaresi et al., 1987). Others have demonstrated extensive labeling of neurons in the non-specific groups of nuclei (e.g. the ventromedial nucleus and the intralaminar nuclei), but only very sparse labeling in specific nuclei (the ventroposterior and ventrolateral nuclei; Ottersen et al., 1983). However, a recent report (Johnson and Burkhalter, 1992) detected extensive retrograde labeling of geniculate neurons following injections of D-[3H]aspartate into the visual cortex, thus demonstrating uptake and transport of D-[3H]aspartate in a population of specific thalamocortical neurons. Retrograde labeling of neurons in several thalamic nuclei, including the mediodorsal nucleus, has also been detected following injections of D-[3H]aspartate into the prefrontal cortex (Pirot et al., 1994). Immunogold studies of Glu content in terminals of thalamocortical neurons have focused on specific sensory projections. Montero (1990) examined collateral terminals of geniculocortical neurons in the cat perigeniculate nucleus, and detected levels of Glu in these terminals that were higher than in their parent cell bodies in the lateral geniculate nucleus. Kharazia and Weinberg (1993, 1994) examined the presence of Glu in anterogradely labeled thalamocortical terminals in the somatic sensory, auditory and visual cortex of rats. Thalamocortical terminals in all these cortical areas were significantly enriched in Glu when compared to the Glu levels detected in dendrites, astrocytes and GABAergic terminals. A significant positive correlation was also noted between the densities of Glu immunogold labeling and synaptic vesicles in thalamocortical terminals but not in GABAergic terminals, a finding which indicates vesicular localization of Glu in thalamocortical terminals. Thus, the available data from D-[3H]aspartate tracing and immunocytochemical studies concur with physiological and pharmacological observations (see e.g.: Tsumoto, 1990; Hicks et al., 1991; McCormick, 1992) in providing strong support for Glu as a transmitter in at least large proportions of the extensive thalamocortical input (Fig. 6). Whether this holds true for all parts of the diverse thalamocortical projections remains to be examined. Similar to thalamocortical terminals, enrichment of Glu is also evident in piriform cortex terminals originating from the olfactory bulb (Hennequet et al., 1998). Zinc-containing terminals (origin unknown) in the cat visual cortex are enriched in Glu but not in GABA (Beaulieu et al., 1992). There are local differences in the density of Glu- and GABA-immunoreactive terminals within the primate striate cortex. Regions in the supragranular layers rich in cytochrome oxidase activity ('puffs') contain higher densities of Glu-immunoreactive terminals and higher ratio of Glu- to GABA-immunoreactive terminals than cytochrome oxidase-sparse regions ('interpuffs'; Nie and Wong-Riley, 1996). The presence of Glu has also been examined in the cerebral cortex of humans (Aas et al., 1993). Slices of macroscopically normal parietal or temporal neocortex were obtained 29

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from patients undergoing surgical treatment for epilepsy or gliomas. Electron microscopical examination of Glu immunogold-labeled sections from this material demonstrated higher levels of Glu in terminals forming asymmetric synaptic contacts than in neuronal cell bodies and glial processes. Glu immunogold findings in the human cerebral cortex thus agree with data from experimental animals.

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Van den Pol AN, Trombley PQ (1993): Glutamate neurons in hypothalamus regulate excitatory transmission. J Neurosci 13:2829-2836. Van der Kooy D, Hattori T (1980): Single subthalamic nucleus neurons project to both the globus pallidus and substantia nigra in rat. J Comp Neurol 192:751-768. Vardi N, Morigiwa K (1997): On cone bipolar cells in rat express the metabotropic receptor mGluR6. Vis Neurosci 14:789-794. Vogel R, Jennemann G, Seitz J, Wiesinger H, Hamprecht B (1998): Mitochondrial malic enzyme: purification from bovine brain, generation of an antiserum, and immunocytochemical localization in neurons of rat brain. J Neurochem 71:844-852. Vollenweider FX, Cu6nod M, Do KQ (1990): Effect of climbing fiber deprivation on release of endogenous aspartate, glutamate, and homocysteate in slices of rat cerebellar hemispheres and vermis. J Neurochem 54:1533-1540. Voogd J, Jaarsma D, Marani E (1996): The cerebellum: chemoarchitecture and anatomy. In: Swanson LW, Bj6rklund A, H6kfelt T (Eds), Handbook of Chemical Neuroanatomy, Vol 12: Integrated Systems of the CNS, Part IlL Amsterdam: Elsevier, pp 1-369. Vyas S, Bradford HF (1987): Co-release of acetylcholine, glutamate and taurine from synaptosomes of Torpedo electric organ. Neurosci Lett 82:58-64. Waelsch H, Berl S, Rossi CA, Clarke DD, Purpura DP (1964): Quantitative aspects of CO2 fixation in mammalian brain in vivo. J Neurochem 11:717-728. Waerhaug O, Ottersen OP (1993): Demonstration of glutamate-like immunoreactivity at rat neuromuscular j unctions by quantitative electron microscopic immunocytochemistry. Anat Embryol (Berl) 188:501-513. Walaas I, Fonnum F (1980): Biochemical evidence for glutamate as a transmitter in hippocampal efferents to the basal forebrain and hypothalamus in rat brain. Neuroscience 5:1691-1698. Walberg F, Ottersen OP (1992): Neuroactive amino acids in the area postrema. An immunocytochemical investigation in rat with some observations in cat and monkey (Macaca fascicularis). Anat Embryol 185:529545. Wanaka A, Shiotani Y, Kiyama H, Matsuyama T, Kamada T, Shiosaka S, Tohyama M (1987): Glutamate-like immunoreactive structures in primary sensory neurons in the rat detected by a specific antiserum against glutamate. Exp Brain Res 65:691-694. Ward HW, Thanki CM, Bradford HF (1983): Glutamine and glucose as precursors of transmitter amino acids: ex vivo studies. J Neurochem 40:855-860. Watkins JC (1986): Twenty-five years of excitatory amino acid research. The end of the beginning? In: Roberts PJ, Storm-Mathisen J, Bradford HF (Eds), Excitatory Amino Acids. London: Macmillan, pp 1-39. Westergaard N, Varming T, Peng L, Sonnewald U, Hertz L, Schousboe A (1993): Uptake, release, and metabolism of alanine in neurons and astrocytes in primary cultures. J Neurosci Res 35:540-545. Westlund KN, Carlton SM, Zhang D, Willis WD (1992): Glutamate-immunoreactive terminals synapse on primate spinothalamic tract cells. J Comp Neurol 322:519-527. Whitesell RR, Ward M, McCall AL, Granner DK, May JM (1995): Coupled glucose transport and metabolism in cultured neuronal cells: determination of the rate-limiting step. J Cereb Blood Flow Metab 15:814-826. Wiklund L, Toggenburger G, Cu6nod M (1984): Selective retrograde labelling of the rat olivocerebellar climbing fiber system with D-[3H]asparate. Neuroscience 13:441-468. Willis WD, Coggeshall RE (1991): Sensory Mechanisms of the Spinal Cord, 2nd ed. New York: Plenum Press. Wilson CJ (1986): Postsynaptic potentials evoked in spiny neostriatal projection neurons by stimulation of ipsilateral and contralateral neocortex. Brain Res 367:201-213. Winter HC, Ueda T (1993): Glutamate uptake system in the presynaptic vesicle: glutamic acid analogs as inhibitors and alternate substrates. Neurochem Res 18:79-85. Yang H-W, Appenteng K, Batten TFC (1997): Ultrastructural subtypes of glutamate-immunoreactive terminals on rat trigeminal motoneurons and their relationships with GABA-immunoreactive terminals. Exp Brain Res 114:99-116. Yoshida M, Teramura M, Sakai M, Karasawa N, Nagatsu T, Nagatsu ! (1987): Immunohistochemical visualization of glutamate- and aspartate-containing nerve terminal pools in the rat limbic structures. Brain Res 410:169-173. Young AB, Oster-Granite ML, Herndon RM, Snyder SH (1974): Glutamic acid: selective depletion by viral induced granule cell loss in hamster cerebellum. Brain Res 73:1-13. Yu ACH, Drejer J, Hertz L, Schousboe A (1983): Pyruvate carboxylase activity in primary cultures of astrocytes and neurons. J Neurochem 41:1484-1487. Yudkoff M (1997): Brain metabolism of branched-chain amino acids. Glia 21:92-98.

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Zeitschel U, Bigl M, Eschrich K, Bigl V (1996): Cellular distribution of 6-phosphofructo-l-kinase isoenzymes in rat brain. J Neurochem 67:2573-2580. Zhang N, Ottersen OP (1992): Differential cellular distribution of two sulphur-containing amino acids in rat cerebellum. An immunocytochemical investigation using antisera to taurine and homocysteic acid. Exp Brain Res 90:11-20. Zhang N, Walberg F, Laake JH, Meldrum BS, Ottersen OP (1990): Aspartate-like and glutamate-like immunoreactivities in the inferior olive and climbing fibre system: a light microscopic and semiquantitative electron microscopic study in rat and baboon (Papio anubis). Neuroscience 38:61-80. Zielke HR, Collins RM, Baab PJ, Huang Y, Zielke CL, Tildon JT (1998): Compartmentation of [14C]glutamate and [14C]glutamine oxidative metabolism in the rat hippocampus as determined by microdialysis. J Neurochem 71:1315-1330.

44

CHAPTER II

neurochemical evidence for a Aspartate transmitter role V. GUNDERSEN AND J. STORM-MATHISEN

1. INTRODUCTION Glutamate (Glu) fulfills most criteria for classification as a neurotransmitter and is regarded as a transmitter (see Chapters I and X) at most excitatory synapses in the mammalian brain. The question of whether aspartate (Asp) serves such a function is still controversial. To have a transmitter role, Asp should be localized in nerve terminals and released by regulated exocytosis (i.e. in a CaZ+-dependent and clostridium toxin sensitive manner) from these terminals. After release, Asp should be able to produce a neuronal response through activation of specific receptors. The first cue that Asp could be a neurotransmitter came from work by Curtis and co-workers in the beginning of the nineteensixties. After application of Asp to central nervous system neurons they could record membrane depolarization (Curtis and Watkins, 1960). Asp has later been shown to activate neurons through the N-methyl-D-aspartate (NMDA) type of Glu receptor (NMDA receptor), whereas it does not seem to have a significant effect on other types of ionotropic Glu receptor (Patneau and Mayer, 1990; Curras and Dingledine, 1992). The question of whether Asp may stimulate the different metabotropic Glu receptors is not resolved. NMDA receptors are formed by two main classes of subunits, NR1 and NR2 (Monyer et al., 1992; Nakanishi, 1992). Although NR1 is essential for functional NMDA receptors (Forrest et al., 1994), this receptor subtype by itself produces only a small response when activated (Moriyoshi et al., 1991). Co-expression of NR1 and one of the NR2 subunits gives larger responses (Kutsuwada et al., 1992; Meguro et al., 1992; Monyer et al., 1992).

2. IS ASPARTATE L O C A L I Z E D IN NERVE TERMINALS?

Like Glu, Asp plays an important role in general cell metabolism and in the synthesis of proteins. Thus, it can be difficult to identify the transmitter pool of Asp and Glu in the brain. By immunocytochemical methods it is possible to visualize the Asp and Glu content separately in different neuronal compartments. One would expect that a substance that has a transmitter role in the brain is localized in nerve endings, which is the site of transmitter release, rather than in neuronal cell bodies and dendrites. Electron microscopical immunogold studies have shown that Glu is enriched in nerve terminals in several excitatory pathways in the brain (Somogyi et al., 1986; Bramham et al., 1990; Maxwell et al., 1990, 1993; Broman and Ottersen, 1992; Walberg and Ottersen, 1992; Handbook of Chemical Neuroanatomy, Vol. 18: Glutamate O.P. Ottersen and J. Storm-Mathisen, editors (~ 2000 Elsevier Science B.V. All rights reserved.

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Broman et al., 1993; Castel et al., 1993; Rinvik and Ottersen, 1993; Zhang and Ottersen, 1993; Davanger et al., 1994; Ericson et al., 1995; Hackney et al., 1996). The immunocytochemical distribution of Asp in perfusion-fixed brain tissue shows conflicting results. Some studies have not detected Asp in nerve endings (Maxwell et al., 1990; Zhang et al., 1990; Ji et al., 1991; Montero, 1994), whereas others have found evidence for such a localization (Merighi et al., 1991; Tracey et al., 1991; Van den Pol, 1991; Usami and Ottersen, 1996). These variable results could be due to differences between the different neuronal pathways studied, or to the low average level of free endogenous Asp in brain tissue, about 1/5 of that of Glu (Nadler et al., 1978), together with a low labeling efficiency of Asp due to the preparation procedure used for embedding the tissue. The latter can be improved by substituting uranyl acetate for osmium tetroxide (Gundersen et al., 1993; Zhang et al., 1994). Since hypoglycemia is known to increase the brain levels of Asp (Butterworth et al., 1982; Engelsen and Fonnum, 1983), one way to overcome the problems with low Asp immunosignals in perfusion-fixed tissue is to combine induction of hypoglycemia in the brain before aldehyde perfusion and treatment of the fixed tissue with uranyl acetate (rather than osmium tetroxide) before embedding. In hippocampi prepared in this way the level of Asp immunogold particles in excitatory nerve terminals was by far higher than in neuronal cell bodies and dendrites (Gundersen et al., 1998; V. Gundersen, unpublished results). Even stronger Asp immunogold signals may be achieved by using tissue embedded in methacrylate resin after freeze-substitution (Usami and Ottersen, 1996; Gundersen et al., 1998). Another way of demonstrating Asp in nerve terminals is to use the brain slice preparation. Hippocampal brain slices seem to show the remarkable property of visualizing the transmitter pool of excitatory amino acids at the expense of the metabolic pool, inasmuch as in such preparations glutamate immunoreactivity is mainly found in nerve endings of excitatory pathways compared to other tissue compartments (Ottersen et al., 1990). By light microscopic immunocytochemistry we have shown that in hippocampal slices Asp, like Glu, is localized in nerve ending-like dots corresponding to the terminal fields of excitatory afferents (Gundersen et al., 1991) (Fig. 1). In slices treated with uranyl acetate before embedding, electron microscopy revealed that these stained dots represent Asp labeling of excitatory nerve terminals (Gundersen et al., 1998). The reason why hippocampal slices fixed by immersion show increased intraterminal Asp content is unclear. Nor seems this a property of all types of brain slice preparations, since in brain stem slices of the dorsal cochlear nucleus, dendrites rather than nerve terminals were immunolabeled with Asp (Osen et al., 1995). It may be that the preparation of slices from the hippocampus causes a flushing away of Asp from cell compartments in which Asp is not protected in synaptic vesicles. This may explain the discrepancy in nerve terminal localization of Asp between hippocampal and dorsal cochlear nucleus slices (see Section 3). During brain slice preparation, Asp may also be eluted from neuronal compartments with low levels of Asp/Glu uptake sites. In favor of this notion is that the hippocampal pattern of Asp immunostaining in the slice preparation resembles the anatomical pattern of uptake sites for Asp/Glu (Storm-Mathisen and Wold, 1981; Taxt and Storm-Mathisen, 1984; Gundersen et al., 1993). Interestingly, the slice staining pattern also resembles the pattern of Asp staining of hypoglycemic hippocampus fixed by perfusion. Indeed, there may be a relative deficit of glucose within the slice tissue. In hypoglycemia, Asp is probably produced from Glu via the aspartate aminotransferase reaction due to accumulation of oxaloacetic acid in the Krebs' cycle (see Engelsen and Fonnum, 1983). Therefore, Asp may rather be synthesized de novo both in the slices and in the perfusion-fixed hypoglycemic hippocampus. Of particular interest concerning de novo synthesis is that in hippocampal slices slightly depolarized in the presence of glutamine (which is a precursor 46

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Fig. 1. Asp-like and Glu-like immunoreactivities in CA1 and area dentata of hippocampus from slices of the hippocampal formation. The slices were fixed by immersion after incubation at physiological K + concentration (5 mM). The photomicrograps show that Asp and Glu are localized with a zonal pattern, corresponding to the distribution of excitatory afferents. Symbols: R, M, layers of hippocampus (radiatum and lacunosum moleculare). Mi, G, layers of area dentata (inner moleculare and granulare). Asterisks mark the obliterated fissura hippocampi. Scale bar -- 100 ~m.

for transmitter Glu) the Asp content in excitatory nerve terminals is increased relative to the resting situation (Gundersen et al., 1991, 1998) (Fig. 2). It has been suggested on the basis of denervation experiments that terminals which synaptically release an excitatory amino acid, have a high turnover of this amino acid (Engelsen and Fonnum, 1983). Thus, the Asp/Glu releasing terminals (see below) may be the ones that are enriched with Asp in the slice preparation and in hypoglycemia in vivo.

3. IS ASPARTATE RELEASED BY EXOCYTOSIS FROM NERVE ENDINGS?

Release of Asp and Glu from nerve terminals may occur through exocytosis of vesicular content or through reversal of plasma membrane transporter proteins for Asp and Glu. Alternatively, Asp could escape from nerve terminals through this transporter system by exchange with synaptically released Glu. Such transporters (see Chapter VIII), besides localization on glial membranes (see Danbolt, 1994), are situated on excitatory nerve terminals (Gundersen et al., 1993, 1996) and they transport Asp and Glu with similar affinities (Balcar and Johnston, 1972; Arriza et al., 1994). In favor of an exocytotic release mechanism 47

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for Asp would be demonstration of sensitivity to factors that inhibit exocytosis, i.e. removal of extracellular C a 2+ or addition of toxins specifically interrupting fusion of synaptic vesicles with the plasma membrane (i.e. Clostridium tetani and C. botulinum toxins (Schiavo et

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al., 1992)). Such a demonstration makes release of Asp through reversal of the transporter proteins less likely, whereas the exchange mechanism cannot be ruled out. One way of inhibiting exchange via the Asp/Glu transporter proteins is to use dihydrokainic acid. This substance is a non-transported inhibitor of GLT1 (Arriza et al., 1994) which is a glial subtype of Asp/Glu transporters. Dihydrokainic acid also inhibits Asp/Glu uptake in hippocampal excitatory nerve terminals (V. Gundersen and Y. Dehnes, unpublished observation). 3.1. RELEASE FROM SYNAPTOSOMES Concerning Ca2+-dependence, release studies using synaptosomes have shown conflicting results. Cortical (Wilkinson and Nicholls, 1989; McMahon and Nicholls, 1990; but see McMahon et al., 1992) and cerebellar synaptosomes (Levi and Gallo, 1981; Levi et al., 1982) have failed to show CaZ+-dependent release of Asp. Thus, the literature has mostly concluded that membrane depolarization-induced release of Asp is CaZ+-independent and that it occurs through reversal of the plasma membrane transporters for Asp/Glu (see Nicholls and Attwell, 1990). However, others have reported CaZ+-dependent release of Asp from both cortical (Pende et al., 1993) and cerebellar (Maura et al., 1991) synaptosomes, as well as from corpus striatum synaptosomes (Maura et al., 1989) using K+-induced membrane depolarization. Recently, Zhou et al. (1995) demonstrated that K + as well as 4-aminopytidine, the latter giving an action potential-like depolarization (Nicholls, 1993), could evoke of CaZ+-dependent Asp release from synaptosomes made from CA1 hippocampal slices. Furthermore, they used an experimental set up in which reuptake was neutralized by fast superfusion and showed that Asp was still released in a Ca2+-dependent manner. Using a similar perfusion assay Breukel et al. (1997) demonstrated that arachidonic acid enhanced K+-induced efflux of both Asp and Glu from hippocampus. Thus, in synaptosomes there is evidence that Asp is neither released due to reversal of the plasma membrane transporters for excitatory amino acids nor to exchange with exocytotically released Glu via the uptake system. This points to an exocytotic release mechanism for Asp. In line with this, tetanus toxin could block CaZ+-dependent release of Asp from cortical synaptosomes (McMahon et al., 1992). Also in cultured neuroendocrine pinealocytes K+-induced release of Asp was both CaZ+-dependent and sensitive to botulinum toxin (Yatsushiro et al., 1997). To rule out involvement of exchange mechanisms in the K+-induced release of Asp the authors used dihydrokainate, which is a non-transported inhibitor (see above) of the excitatory amino acid carrier (GLT1) found in the pinealocytes (Yamada et al., 1997). This treatment did not affect the K+-induced release, but inhibited Na+-dependent sequestration of Asp into the pinealocytes. In addition they showed that the pinealocytes, which contain both the L-form the D-form of Asp, released only L-Asp upon membrane depolarization. This makes it further

<-

Fig.

2. Light micrographs showing the distribution of Asp-like immunoreactivity in slices of hippocampus. (A) Slice incubated in 5 mM K +. (B) 40 mM K +. (C) 55 mM K +. (D) 5 mM K + plus 0.5 mM glutamine. (E) 40 mM K + plus 0.5 mM glutamine. (F) 55 mM K + plus 0.5 mM glutamine. Note the gradual disappearance of the zonal staining pattern ('nerve terminal-like staining pattern') and appearance of Asp staining in structures representing glial cells at high K + concentrations and that these changes are inhibited by glutamine. When the slices are moderately depolarized (40 mM K +) in the presence of glutamine, the staining intensity of Asp in the terminal zones of excitatory afferents is enhanced (E). Symbols: O, P, R, M, layers of hippocampus (oriens, pyramidale, radiatum, lacunosum moleculare). Triangles mark the inner molecular layer of area dentata. G, granule layer of area dentata. Asterisks mark the obliterated fissura hippocampi. Scale bar - 100 [~m. (Modified from Gundersen et al., 1991.)

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unlikely that Asp is released through the plasma membrane transporters, since the latter carry L- and D-Asp with similar affinities (Balcar and Johnston, 1972). 3.2. RELEASE FROM BRAIN SLICES In a more intact in vitro preparation, i.e. the hippocampal brain slice, there is a large body of data showing that the release of Asp upon membrane depolarization is CaZ+-dependent (Nadler et al., 1976, 1990; Burke and Nadler, 1988; Szerb, 1988; Martin et al., 1991; Roisin et al., 1991; Fleck et al., 1993; Zhou et al., 1995). In addition, there is evidence for a CaZ+-dependent release of Asp in cerebellar slices (Flint et al., 1981; Toggenburger et al., 1983; Sekiguchi et al., 1986; Vollenweider et al., 1990; Maura et al., 1991), striatal slices (Reubi et al., 1980; Umeda and Sumi, 1989; Kimura et al., 1995) and spinal cord slices (Kangrga and Randic, 1990). 3.3. RELEASE FROM THE INTACT BRAIN Also in the intact brain push pull cannula and microdialysis experiments have demonstrated K+-evoked CaZ+-dependent Asp release from the striatum (Girault et al., 1986; Paulsen and Fonnum, 1989; but see Zuiderwijk et al., 1996). Recently Lada et al. (1998) demonstrated Ca2+-dependent release of Asp from the striatum after electrical stimulation of the prefrontal cortex. Experiments interfering with the exocytotic machinery have also been performed in the intact striatum. Analysis of microdialysates (Herrera-Marschitz et al., 1996) has shown that extracellular Asp levels were decreased during K+-induced depolarization after treatment with alpha-latrotoxin, which triggers sustained exocytosis (Henkel and Sankaranarayanan, 1999). This is presumably because of depletion of the vesicular content of Asp before the stimulated release. However, care should be taken in interpreting this result, because alpha-latrotoxin may release both vesicular and cytoplasmic pools of amino acids (McMahon et al., 1990). Nonetheless, most experiments in the intact brain, using both direct chemical depolarization of the tissue and stimulation of pathways, show Asp release consistent with exocytosis. 3.4. RELEASE BY HETEROEXCHANGE? One possible concern is that Ca2+-dependent release of Asp in brain slices and in the intact brain could reflect exchange of intracellular Asp with extracellular Glu that has been released by exocytosis (see above). Opposing this idea is the demonstration of independent regulation of Asp and Glu release. Since plasma membrane Asp/Glu transporters have similar affinities for Asp and Glu, it seems unlikely that separate overflow of Asp and Glu is due to regulation at the level of the transporters. However, this cannot be completely excluded, as artificially mutated GLT1 has shown discrimination of Asp and Glu (Zhang et al., 1994). In the hippocampal slice, activation of NMDA-receptors enhances the release of both Asp and Glu, whereas activation of non-NMDA-receptors selectively depressed the K+-evoked release of Asp (Martin et al., 1991; for synaptosomes see Zhou et al., 1995). In addition, a 5-1ipoxygenase product selectively enhances Asp release and a cyclooxygenase product selectively depresses glutamate release (Peterson et al., 1995). Interestingly, using a cochlear in vitro preparation, Jfiger et al. (1998) could demonstrate that increasing the intensity of sound stimuli caused an enhanced release of Asp, whereas an evoked efflux of Glu was seen only at the highest sound intensity. During conditions with reduced glucose concentration in the extracellular fluid, K+-induced membrane depolarization causes the ratio of Asp to Glu, released CaZ+-dependently from hippocampal slices, to increase (Szerb and O'Regan, 1987; 50

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t

Fig. 3. Electron micrographs showing gold particles signaling L-Asp in stratum radiatum CA1 of hippocampal slices incubated at 55 mM K + (A) and 55 mM K + with low Ca 2+/high Mg 2+ (B) for 1 h before fixation. In A note that the terminal with asymmetrical junction (at) on a spine (s) is weakly labeled, whereas the similar type of terminal in B is strongly labeled. The glial profile (g) in A is strongly immunopositive for L-Asp. (The glial mitochondrion (m) is heavily labeled, other mitochondria not.) In B also note the contrast in labeling between the terminals (at) and the spines (s). Inset in A, a higher power photomicrograph of a part of the terminal (at) in A showing individual synaptic vesicles with diameters of about 20-60 nm (arrowheads), which is in the same range as in this type of terminal in vivo (Harris and Sultan, 1995, i.e. the terminal still has synaptic vesicles, but these are depleted of Asp). Asterisks, synaptic cleft. Scale bars: 200 nm in A and B; 100 nm in inset. (From Gundersen et al., 1998.)

Fleck et al., 1993). This result was also obtained under conditions mimicking high neuronal activity (Szerb, 1988). Also in the intact striatum using the microdialysis technique, a selective increase in Asp release after chemical depolarization (with GABA antagonists or NMDA) of the neocortex has been demonstrated (Palmer et al., 1989). Such selectivity suggests that Asp is released independently of Glu both in brain slices and the intact brain. Furthermore, in the intact striatum dihydrokainic acid increased the extracellular level of both Asp and Glu during K + depolarization (Herrera-Marschitz et al., 1996). If heteroexchange was responsible for the depolarization-induced increase in extracellular Asp one would expect the Asp levels to be decreased by dihydrokainic acid (but see Fallgren and Paulsen, 1996). 3.5. IMMUNOCYTOCHEMICAL OBSERVATIONS Adding to this biochemical and electrophysiological evidence for exocytotic Asp release, are observations by light microscopical immunocytochemistry showing that K + depolarization of hippocampal slices induces a depletion of Asp immunoreactive nerve ending-like dots in the terminal areas of excitatory fibers (Gundersen et al., 1991) (Fig. 2). Electron microscopic investigation demonstrated that excitatory nerve terminals, which were labeled with Asp immunogold particles in slices that had been incubated at a physiological K + concentration, were depleted of their Asp immmunoreactivity during K+-induced membrane depolarization. This change was inhibited by low-Ca 2+ conditions (Fig. 3) and by tetanus toxin (Gundersen et al., 1998).

4. IS ASPARTATE LOCALIZED IN SYNAPTIC VESICLES? Essential as evidence for exocytotic Asp release would be a demonstration of Asp in synaptic vesicles and of a vesicular uptake system for Asp. In contrast to Glu, no study has so 51

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Fig. 4. Electron micrographs of L-Asp-LI and L-Glu-LI in hippocampus CA1 from a hypoglycemic rat subjected to perfusion fixation. The tissue was treated with uranyl acetate before embedding in epoxy resin. The figure shows accumulation of immunoreactivities over synaptic vesicle clusters (sv) versus over cytoplasmic matrix (cm) in terminals making asymmetrical synapses on spines (s). Broken lines mark the boundary between the vesicle-rich and vesicle-poor parts of the terminals. Scale bar = 0.2 ~m. (Modified from Gundersen et al., 1998.)

far shown uptake (Naito and Ueda, 1983; Maycox et al., 1988; Fykse et al., 1992) or content (Burger et al., 1991) of Asp in isolated synaptic vesicles from the brain. On the other hand, Cousin and Nicholls (1997) could give indirect evidence for synaptic vesicle uptake of an excitatory amino acid other than Glu in cultured cerebellar granule cells. They showed that after preloading the neurons with exogenous D-Asp this 'false transmitter' was released in a strictly exocytotic manner following electrical stimulation (release of D-Asp was inter alia blocked by bafilomycin A1, which disrupts the transmembrane vesicular proton-electrochemical gradient). In line with this, we could, by immunoelectron microscopy in the hypoglycemic hippocampus fixed by perfusion, demonstrate that Asp was concentrated over synaptic vesicle-rich parts of excitatory nerve terminals relative to over vesicle-poor parts (Gundersen et al., 1998) (Fig. 4). The labeling ratio of Asp between these compartments was comparable to that of Glu and significantly higher than those of glutamine and taurine. Also in excitatory terminals from freeze-substitution embedded normoglycemic hippocampus Asp immunogold particles were located much closer to the center of synaptic vesicles than glutamine immunogold particles, suggesting that differences in fixation efficiency between the vesicle cluster and cytosolic matrix cannot explain the vesicular Asp accumulation. Also in favor of a synaptic vesicle localization of Asp, is that Asp immunoreactivity was found to be co-localized with synaptophysin, a synaptic vesicle protein, in neuroendocrine pinealocyte microvesicles shown to release Asp in an exocytotic manner (Yatsushiro et al., 1997). Thus, synaptic vesicles in situ seem to be able to accumulate Asp. This ability seems to be easily lost on isolation of the vesicles.

5. IS ASPARTATE RELEASED FROM A SEPARATE POOL OF NERVE ENDINGS?

The biochemical data from the hippocampus indicating that Asp is released independently of Glu suggest that there are either separate Asp- and Glu-containing terminals, or that Glu and Asp are located in the same terminal but in different synaptic vesicles that can be independently regulated. Our immunogold results from hippocampal slices and from hypoglycemic perfusion-fixed hippocampi showed that Asp and Glu are localized in the same excitatory nerve terminals (Gundersen et al., 1998) (Fig. 5). We did not find evidence for a separate pool of Asp-ergic nerve terminals. Since the immunogold method did not have high 52

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Fig. 5. Neighboring ultrathin sections from hypoglycemic CA1 hippocampus (same block as in Fig. 4) showing co-localization of L-Asp (A) and L-Glu (B) immunogold particles in a terminal (atl) with asymmetrical synaptic specialization (arrows) on a spine (s). Note that the terminal has high levels of both immunoreactivities, whereas the spines and dendritic shafts (d) are very weakly labeled. Scale bar = 0.2 I~m. (Modified from Gundersen et al., 1998.)

enough resolution to address the question of whether the amino acids were located in the same or different pools of synaptic vesicles, we can at present only speculate about this matter. It should be mentioned that in the spinal cord, Merighi et al. (1991) and Valtschanoff et al. (1993) found evidence for different pools of Asp- and Glu-containing excitatory nerve terminals.

6. THE ROLE OF THE RELEASED ASPARTATE As discussed above, there is evidence suggesting that Asp is localized and synaptically released from the terminals of several excitatory nerve terminal systems in the brain. Especially interesting is that when hippocampal slices are stimulated in the presence of glutamine, the content of Asp in excitatory nerve terminals increases (Gundersen et al., 1991, 1998) and that increased neuronal activity results in an enhanced release of Asp from hippocampal slices (Szerb, 1988). Hypoglycemic conditions may mimic conditions under high synaptic activity, inasmuch as the ratio between energy demand and supply is increased. Under hypoglycemia there is also an enrichment of Asp in excitatory nerve terminals (Gundersen et al., 1998). It may therefore be that during increased synaptic activity, excitatory nerve terminals can build up and release increased amounts of Asp. One criterion for classifying a substance as a neurotransmitter is that it should activate postsynaptic receptors after release from terminals. Fleck et al. (1993) found that endogenous released Asp brought about a rapid postsynaptic NMDA-receptor-mediated response after stimulation of the Schaffer collateral fibers in hippocampal slices when glucose concentration was reduced, indicating a functional role of the released Asp. By itself Asp can hardly mediate fast synaptic transmission, since non-NMDA receptors are responsible for most of the postsynaptic current at central excitatory synapses and, as discussed above, these appear insensitive to Asp. However, Asp could cause a shift in the postsynaptic excitatory response towards NMDA receptor activation, a mechanism that would be important for several aspects of excitatory neurotransmission. Asp may therefore be involved in the induction of the NMDA-dependent type of long-term potentiation. Indeed, Asp is released during the induction of LTP in CA1 of the hippocampus 53

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(Bliss et al., 1986). Furthermore, release of Asp may underlie the pure NMDA receptor response known to occur at synapses where AMPA receptors are also thought to be present (Liao et al., 1995). Alternatively, Asp stimulation of NMDA receptors may trigger metabolic events in the brain. Activation of NMDA receptors induce production of nitric oxide (NO) through stimulation of NO synthase (Garthwaite, 1991). This enzyme is located postsynaptically at excitatory synapses (Aoki et al., 1993). It is well known that NO dilates cerebral blood vessels and it has in fact been demonstrated that activation of NMDA receptors during high synaptic activity causes a local vasodilatation which is mediated by NO production (Faraci and Breese, 1993). Thus, it is tempting to speculate that Asp could be released to increase the flow of oxygen and glucose to active synapses. In addition, Asp has been implicated in the generation of seizures in epileptic (EL) mice, which are regarded as a model of human temporal lobe epilepsy. Flavin and Seyfried (1994) showed that the CaZ+-dependent K+-induced release of Asp was higher in hippocampal slices made from EL mice before seizures occurred than in slices from genetically related non-seizure prone mice. This was not the case for Glu or GABA. It remains to be elucidated whether NMDA receptor responses are involved in the seizure generation in EL mice.

7. PUTATIVE ASPARTATERGIC NEURONAL PATHWAYS 7.1. THE HIPPOCAMPAL FORMATION Based on the following data it may be concluded that Asp is a transmitter candidate in the hippocampal formation at excitatory nerve terminals of the Schaffer collateral-commissural projection. (1) Biochemical data from denervated hippocampal slices and hippocampal slices, in which CA3 pyramidal neurons were destroyed with kainic acid, have shown that Asp is released from this pathway in a Ca2+-dependent manner (Nadler et al., 1976; Burke and Nadler, 1988). (2) Immunocytochemical observations from hippocampal slices and perfusion-fixed hippocampus have suggested that Asp is located in synaptic vesicles in excitatory nerve endings of the Schaffer collateral-commissural synapses and that Asp is depleted in a manner consistent with exocytosis from these terminals (Gundersen et al., 1998). (3) Electrophysiological experiments have demonstrated that stimulation of the Schaffer collaterals caused an Asp-mediated NMDA response (Fleck et al., 1993). (4) NMDA receptors have been localized by immunocytochemistry to the postsynaptic membrane of Schaffer collateral-commissural synapses (Petralia et al., 1994a,b; Petralia et al., 1999; Takumi et al., 1999). In addition, we have shown that Asp is localized in and exocytotically depletable from nerve terminals of the other excitatory pathways in the hippocampal formation (Gundersen et al., 1991, 1998). Interestingly, NMDA receptor labeling of hippocampus has shown particular strong staining of the termination zones of the CA4 mossy cell axons in the inner third of dentate gyms (Petralia et al., 1994a,b). This is an area with a strong Asp staining, especially after stimulation of the slices in the presence of glutamine (Gundersen et al., 1991). It should also be mentioned that commissurotomy reduced the CaZ+-dependent release of Asp, but not Glu, in slices of dentate gyrus (Nadler et al., 1976), further substantiating the idea that synaptic transmission in this part of the hippocampal formation is mediated by Asp. 54

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7.2. STRIATUM Biochemically, there is evidence for Ca2+-dependent release of Asp after chemical depolarization in striatum (Reubi et al., 1980; Girault et al., 1986; Paulsen and Fonnum, 1989; Umeda and Sumi, 1989; Kimura et al., 1995). This is the case also after electric stimulation of the corticostriatal pathway (Lada et al., 1998). Light-microscopical observations have concluded that Asp immunoreactivity may be localized in nerve terminals of striatal interneurons (Snyder et al., 1993; Pettersson et al., 1996) and in most cortical neurons projecting to the striatum (Bellomo et al., 1998). However, to our knowledge, there are at present no electron microscopical observations showing the distribution of Asp immunoreactivity between the different nerve endings in the striatum. Thus, a conclusion concerning Asp-ergic synaptic transmission in striatum, including corticostriatal nerve endings, should await results of such studies. 7.3. CEREBELLAR CORTEX The transmitter role of Asp in the major excitatory nerve fiber systems in the cerebellar cortex, which consist of the climbing and parallel fiber inputs to Purkinje cells and the mossy fiber projection to granule cells, is unclear. Especially its role at the olivocerebellar climbing fiber synapses has been extensively debated (see Zhang and Ottersen, 1993). Biochemically, there is evidence for Asp-ergic synaptic transmission in the cerebellar cortex. Investigations of cerebellar slices following destruction of the climbing fibers have revealed a relatively larger reduction in the CaZ+-dependent release of Asp than in that of Glu (26% vs. 14%) (Wiklund et al., 1982; Toggenburger et al., 1983). This result was confirmed in climbing fiber deprived cerebellar slices from the hemisphere but not from vermis (Vollenweider et al., 1990). Based on these observations, Asp was proposed as the excitatory transmitter in cerebellar climbing fibers. However, since the reduction in CaZ+-dependent release in absolute terms was by far higher for Glu than for Asp, and since a huge release of Glu from parallel fibers could mask the effect of a loss of climbing fibers, these experiments indicate that both Asp and Glu might be transmitters at climbing fiber synapses. Cerebellar slice experiments have also suggested that Asp is released CaZ+-dependently from parallel fibers (Flint et al., 1981; Vollenweider et al., 1990). In line with the in vitro data discussed above are immunocytochemical results from cerebellar slices incubated in vitro, showing Asp labeling of climbing fibers at a physiological K + concentration. Incubation in a high K + concentration abolished this labeling (Ottersen and Laake, 1992). Opposing the idea that Asp is a transmitter in the main excitatory fiber systems in the cerebellum, are electron-microscopical immunogold studies of the cerebellar hemisphere fixed by in vivo perfusion. In such an intact cerebellum low densities of Asp immunogold particles have been found in climbing fiber and parallel fiber terminals, as opposed to neuronal cell bodies in the inferior olive (Zhang et al., 1990), as well as in mossy fiber terminals (Ji et al., 1991). The discrepancy in Asp labeling between the in vivo and in vitro fixed tissue could be due to the fact that the slices allow Asp to accumulate in nerve fibers and terminals (see Section 2). It should be mentioned that Campistron et al. (1986) have presented a light micrograph of in vivo fixed cerebellum showing Asp immunoreactivity in structures resembling climbing and parallel fibers. Thus, based on immunocytochemical results, we feel that the question whether Asp has a transmitter role in the excitatory pathways in the cerebellum is not resolved. A conclusion concerning the distribution of Asp immunoreactivity between nerve endings and other neuronal elements in the intact cerebellar cortex should 55

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await results from tissue prepared by methods known to give a strong Asp immunosignal (see Section 2). Another argument against Asp-ergic synaptic transmission in mature cerebellar paralleland climbing-fiber nerve terminals, would be that most studies have shown that NMDA receptor responses are present only early in development in Purkinje cells (Hirano, 1990; Rosenmund et al., 1992; Yuzaki et al., 1996a; Hausser and Roth, 1997). However, Sekiguchi et al. (1986) have demonstrated NMDA-induced currents in Purkinje cells in adult cerebellar slices. A functional NMDA receptor is thought to consist of NR1 and a combination of the different subtypes of NR2 (see Section 1). Indeed, NR1 (Moriyoshi et al., 1991; Akazawa et al., 1994; Monyer et al., 1994; Petralia et al., 1994a) as well as NR2 (Akazawa et al., 1994; Petralia et al., 1994b, but see Monyer et al., 1994) mRNA and protein seem to be expressed in mature Purkinje cells. In addition, Yuzaki et al. (1996b) demonstrated that Asp could selectively activate a Ca 2+ response in Purkinje cells. However, the mechanism responsible for this response has not yet been identified. 7.4. SPINAL CORD In the case of primary afferents to the spinal dorsal horn there are conflicting results as to whether the terminals of these afferents contain Asp. Maxwell et al. (1990) found no evidence for enrichment of Asp immunolabeling in primary afferent terminals in the cat, but in the rat such evidence was found by Merighi et al. (1991) and Tracey et al. (1991). In addition, the presence of Asp immunoreactivity in rat dorsal root axons has been reported (Westlund et al., 1989). In line with the latter studies, Asp has been shown to be released in a CaZ+-dependent manner from the spinal dorsal horn slice preparation after electrical stimulation of the dorsal root (Kangrga and Randic, 1990). Furthermore, NMDA responses could be activated when primary afferents were electrically stimulated (Gerber and Randic, 1989; Gerber et al., 1991). The NMDA receptor protein has been detected in the terminal area of the primary afferents (Yung, 1998). Taken together, the experimental data from the spinal dorsal horn seem to support a transmitter candidacy of Asp in the primary afferent fibers. By combining electron-microscopical immunocytochemistry with anterograde tracing of corticospinal terminals, Asp was found to be localized in a subpopulation of these terminals (Valtschanoff et al., 1993). However, Asp labeling was much weaker in the identified nerve terminals than in postsynaptic dendritic spines, possibly indicating a metabolic role of Asp. Exocytotic release of Asp should therefore be demonstrated before considering Asp as a transmitter in the corticospinal system. 7.5. AUDITIVE SYSTEMS By using tissue prepared by the freeze-substitution technique, Asp immunogold particles were found in presynaptic terminals of hair cells in the cochlea (Usami and Ottersen, 1996, but see Liu, 1997). Recently Jfiger et al. (1998) gave evidence for release of Asp from a cochlear in vitro preparation after physiological sound stimulation, implying that Asp should be regarded as a transmitter candidate at hair cell synapses. It is interesting that mRNA coding for the NMDA receptor has been localized to spiral ganglion cells (Safieddine and Eybalin, 1992; Kuriyama et al., 1993). Concerning the central auditive pathways, the role of Asp is unresolved. However, based on the immunogold study of Osen et al. (1995) Asp seems not to contribute to synaptic transmission at the parallel fiber synapse in the dorsal cochlear nucleus. 56

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7.6. VISUAL SYSTEMS Although Asp immunoreactivity was shown to be present in terminals of photoreceptors and bipolar cells in the retina, the Asp level seemed higher in the parent cell bodies (Jojich and Pourcho, 1996), questioning the transmitter role of Asp in these nerve endings. Also in the further transmission of visual information Asp seems not to be involved, inasmuch as Montero (1994) found only a low density of Asp immunoreactivity in nerve terminals of retinogeniculate and geniculocortico fibers. Again, until these Asp labeling patterns are confirmed in tissue which is prepared to give a strong Asp immunosignal (see Section 2), care should be taken in concluding about the transmitter role of Asp.

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Somogyi P, Halasy K, Somogyi J, Storm-Mathisen J, Ottersen OP (1986): Quantification of immunogold labelling reveals enrichment of glutamate in mossy and parallel fibre terminals in cat cerebellum. Neuroscience 19:10451050. Storm-Mathisen J, Wold JE (1981): In vivo high-affinity uptake and axonal transport of D-[2,3-3H]aspartate in excitatory neurons. Brain Res 230:427-433. Szerb JC (1988): Changes in the relative amounts of aspartate and glutamate released and retained in hippocampal slices during stimulation. J Neurochem 50:219-224. Szerb JC, O'Regan PA (1987): Reversible shifts in the Ca2+-dependent release of aspartate and glutamate from hippocampal slices with changing glucose concentrations. Synapse 1:265-272. Takumi Y, Ramirez-Leon V, Laake R Rinvik E, Ottersen OP (1999): Different modes of expression of AMPA and NMDA receptors in hippocampal synapses. Nat Neurosci 2:618-624. Taxt T, Storm-Mathisen J (1984): Uptake of D-aspartate and L-glutamate in excitatory axon terminals in hippocampus: autoradiographic and biochemical comparison with gamma-aminobutyrate and other amino acids in normal rats and in rats with lesions. Neuroscience 11:79-100. Toggenburger G, Wiklund L, Henke H, Cu6nod M (1983): Release of endogenous and accumulated exogenous amino acids from slices of normal and climbing fibre-deprived rat cerebellar slices. J Neurochem 41:1606-1613. Tracey DJ, De BS, Phend K, Rustioni A (1991): Aspartate-like immunoreactivity in primary afferent neurons. Neuroscience 40:673-686. Umeda Y, Sumi T (1989): Evoked release of endogenous amino acids from rat striatal slices and its modulation. Eur J Pharmacol 163:291-297.

Usami S, Ottersen OP (1996): Aspartate is enriched in sensory cells and subpopulations of non-neuronal cells in the guinea pig inner ear: a quantitative immunoelectron microscopic analysis. Brain Res 742:43-49. Valtschanoff JG, Weinberg RJ, Rustioni A (1993): Amino acid immunoreactivity in corticospinal terminals. Exp Brain Res 93:95-103. Van den Pol AN (1991): Glutamate and aspartate immunoreactivity in hypothalamic presynaptic axons. J Neurosci 11:2087-2101. Vollenweider FX, Cu6nod M, Do KQ (1990): Effect of climbing fiber deprivation on release of endogenous aspartate, glutamate, and homocysteate in slices of rat cerebellar hemispheres and vermis. J Neurochem 54:1533-1540. Walberg F, Ottersen OP (1992): Neuroactive amino acids in the area postrema. An immunocytochemical investigation in rat with some observations in cat and monkey (Macaca fascicularis). Anat Embryol (Berl) 185:529545. Westlund KN, McNeill DL, Patterson JT, Coggeshall RE (1989): Aspartate immunoreactive axons in normal rat L4 dorsal roots. Brain Res 489:347-351. Wiklund L, Toggenburger G, Cu6nod M (1982): Aspartate: possible neurotransmitter in cerebellar climbing fibers. Science 216:78-80.

Wilkinson R, Nicholls DG (1989): Compartmentation of glutamate and aspartate within cerebral-cortical synaptosomes: evidence for a non-cytoplasmic origin for the CaZ+-releasable pool of glutamate. Neurochem Int 15:191-197. Yamada H, Yatsushiro S, Yamamoto A, Hayashi M, Nishi T, Futai M, Yamaguchi A, Moriyama Y (1997): Functional expression of a GLT-1 type Na+-dependent glutamate transporter in rat pinealocytes. J Neurochem 69:1491-1498. Yatsushiro S, Yamada H, Kozaki S, Kumon H, Michibata H, Yamamoto A, Moriyama Y (1997): L-aspartate but not the D form is secreted through microvesicle-mediated exocytosis and is sequestered through Na+-dependent transporter in rat pinealocytes. J Neurochem 69:340-347. Yung KK (1998): Localization of glutamate receptors in dorsal horn of rat spinal cord. Neuroreport 9:1639-1644. Yuzaki M, Forrest D, Verselis LM, Sun SC, Curran T, Connor JA (1996a): Functional NMDA receptors are transiently active and support the survival of Purkinje cells in culture. J Neurosci 16:4651-4661. Yuzaki M, Forrest D, Curran T, Connor JA (1996b): Selective activation of calcium permeability by aspartate in Purkinje cells. Science 273:1112-1114.

61

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Zhang N, Ottersen OP (1993): In search of the identity of the cerebellar climbing fiber transmitter: immunocytochemical studies in rats. Can J Neurol Sci 3:36-42. Zhang N, Walberg F, Laake JH, Meldrum BS, Ottersen OP (1990): Aspartate-like and glutamate-like immunoreactivities in the inferior olive and climbing fibre system: a light microscopic and semiquantitative electron microscopic study in rat and baboon (Papio anubis). Neuroscience 38:61-80. Zhang Y, Pines G, Kanner BI (1994): Histidine 326 is critical for the function of GLT-1, a (Na + + K+ )-coupled glutamate transporter from rat brain. J Biol Chem 269:19573-19577. Zhou M, Peterson CL, Lu YB, Nadler JV (1995): Release of glutamate and aspartate from CA1 synaptosomes: selective modulation of aspartate release by ionotropic glutamate receptor ligands. J Neurochem 64:1556-1566. Zuiderwijk M, Veenstra E, Lopes da Silva FH, Ghijsen WE (1996): Effects of uptake carrier blockers SK and F 89976-A and L-trans-PDC on in vivo release of amino acids in rat hippocampus. Eur J Pharmaco1307:275-282.

62

CHAPTER III

Metabotropic glutamate receptors immunocytochemical and in situ hybridization analyses R. SHIGEMOTO AND N. MIZUNO

1. INTRODUCTION Metabotropic glutamate receptors (mGluRs), which are linked to several intracellular signal transduction mechanisms via G-protein, are implicated in diverse functions of the mammalian central nervous system (CNS). These functions include mediation of slow excitatory (Glaum and Miller, 1992; McCormick and Von Krosigk, 1992; Eaton et al., 1993) and inhibitory (Fiorillo and Williams, 1998) responses, regulation of calcium channels (Swartz and Bean, 1992; Sahara and Westbrook, 1993; Chavis et al., 1994), potassium channels (Charpak et al., 1990; Shirasaki et al., 1994; Gereau and Conn, 1995; Netzeband et al., 1997) and non-selective cation channels (Gu6rineau et al., 1995; Congar et al., 1997), inhibition (Baskys and Malenka, 1991; Desai and Conn, 1991; Schrader and Tasker, 1997) and facilitation (Herrero et al., 1992; Rodrfguez-Moreno et al., 1998) of transmitter release, induction of long-term potentiation (Bortolotto and Collingridge, 1993; O'Connor et al., 1995; Manahan-Vaughan, 1997) and long-term depression (Linden et al., 1991; Kato, 1993; Shigemoto et al., 1993; Bolshakov and Siegelbaum, 1994; Conquet et al., 1994), and regulation of neuronal development (Hensch and Stryker, 1997; Kano et al., 1997; Plenz and Kitai, 1998). Functional diversity of the metabotropic glutamate receptor is reflected in molecular diversity of receptor subtypes. Eight different cDNAs encoding metabotropic glutamate receptor subtypes, termed mGluR1 through mGluR8, have so far been cloned and alternative splice forms have been known to occur for mGluR1 (mGluRla, mGluRlb, mGluRlc, mGluRld, and mGluRlg), mGluR4 (mGluR4a and mGluR4b), mGluR5 (mGluR5a and mGluR5b), mGluR7 (mGluR7a and mGluR7b), and mGluR8 (mGluR8a and mGluR8b) (Flor et al., 1997; Makoff et al., 1997; Mary et al., 1997; Corti et al., 1998; for further review, see Pin and Duvoisin, 1995). The eight mGluR subtypes are classified into three groups according to their amino acid sequence similarities, preferred signal transduction mechanisms, and pharmacological properties: group I mGluRs, mGluR1 and mGluR5, are selectively activated by 3,5-dihydroxyphenylglycine (DHPG, Schoepp et al., 1994) and are coupled to inositol phospholipid hydrolysis (Masu et al., 1991; Abe et al., 1992); group II mGluRs, mGluR2 and mGluR3, are selectively activated by (+)-lS,2S,5R,6S-2-aminobicyclo[3.1.O]hexane-2,6-dicarboxylic acid (LY354740, Monn et al., 1997) and (2S,l'R,2'R,3'R)-2-(2,3-dicarboxycyclopropyl) glycine (DCG-IV, Hayashi et al., 1993) and are coupled to the inhibition of forskolin-stimulated cyclic AMP production

Handbook of Chemical Neuroanatomy, Vol. 18: Glutamate O.E Ottersen and J. Storm-Mathisen, editors (~) 2000 Elsevier Science B.V. All rights reserved.

63

Ch. III

R. Shigemoto and N. Mizuno

(Tanabe et al., 1992, 1993); group III mGluRs, mGluR4, mGluR6, mGluR7 and mGluR8, are selectively activated by L-2-amino-4-phosphonobutyrate (c-AP4) and are coupled to the inhibitory cyclic AMP cascade (Nakajima et al., 1993; Tanabe et al., 1993; Okamoto et al., 1994; Duvoisin et al., 1995; Saugstad et al., 1997; for review, see Pin and Duvoisin, 1995; Conn and Pin, 1997). The second messenger systems for these mGluRs described in initial cloning studies, however, were all examined in receptor cDNA-transfected heterologous cell lines and it is not necessarily clear which G-proteins and effector molecules are utilized in various neuronal cells in vivo. For example, group I mGluRs are indeed coupled to phospholipase C and subsequent production of inositol triphosphates and induces intracellular calcium release in Purkinje cells (Yuzaki and Mikoshiba, 1992; Takechi et al., 1998) and hippocampal CA1 neurons (Frenguelli et al., 1993), but the same receptor subtypes are also coupled to inhibition of voltage-dependent calcium channels in hippocampal neurons without intracellular diffusible messengers (Lester and Jahr, 1990; Swartz and Bean, 1992). In the latter case, activated G-proteins seem to interact directly with the calcium channels in a membrane delimited manner. Group II mGluRs can be coupled to inhibition of cyclic AMP cascade in neuronal and glial cells (Baba et al., 1993; Pr6zeau et al., 1994) but also linked to rapid-onset regulation of various channels including calcium channels (Choi and Lovinger, 1996) and G-protein-coupled inwardly rectifying K + channels (GIRK) (Knoflach and Kemp, 1998) depending on neuronal cell types (for review, see Anwyl, 1999). Except mGluR6-mediated synaptic transmission to ON bipolar cells in the retina (Masu et al., 1995), the group III mGluR-mediated effect is mostly inhibition of neurotransmission (Trombley and Westbrook, 1992; Jane et al., 1994) through suppression of presynaptic voltage-dependent calcium channels (Takahashi et al., 1996). In certain forms of synaptic plasticity, however, group III mGluRs also seem to be involved as reported in the cerebellum (Pekhletski et al., 1996) and basolateral amygdala (Neugebauer et al., 1997). Thus, mGluRs in the three subgroups have a large variety of transduction mechanisms depending on receptor subtypes, cell types in which they are expressed, and effector molecules associating with mGluRs. The fast excitatory neurotransmission in mammalian brain is mainly mediated by ionotropic glutamate receptors localized largely in postsynaptic membrane specialization of glutamatergic synapses. On the other hand, the sites of action of mGluRs are more widely found throughout different membrane compartments of neuronal and glial cells. For example, regulation of transmitter release mediated by mGluRs is reported not only in glutamatergic synapses but also in GABAergic synapses (Desai and Conn, 1991; Stefani et al., 1994; Poncer et al., 1995; Kinoshita et al., 1998; Bradley et al., 1999) and dopaminergic system (Hu et al., 1999). Furthermore, even in the glutamatergic system, activation of mGluRs by synaptically released glutamate often requires high-frequency or repetitive stimulation (Batchelor et al., 1994; Yokoi et al., 1996; Congar et al., 1997; Scanziani et al., 1997; FioriIlo and Williams, 1998), possibly due to extrasynaptic location of mGluRs on dendrites and axons. These situations may also imply signal transmission between different synapses (heterosynaptic interaction) or different pathways mediated by spillover glutamate activating mGluRs remote from glutamate release sites (Ohishi et al., 1994; Wada et al., 1998; Vogt and Nicoll, 1999). To understand the diversified physiological effects of glutamate, it is thus important to know molecular identity of mGluRs expressed in distinct subpopulations of neurons, membrane compartments of neurons they are localized to, and spatial relation between mGluRs and glutamate release sites of identified origins. In this chapter, regional and cellular distribution of eight mGluRs in the mammalian CNS will be first reviewed and then, distinct subcellular localizations of mGluRs in three subgroups and its functional implication will be discussed. 64

Metabotropic glutamate receptors

Ch. III

2. REGIONAL AND CELLULAR LOCALIZATION OF METABOTROPIC GLUTAMATE RECEPTORS 2.1. AN OVERVIEW Many studies by in situ hybridization histochemistry and immunohistochemistry have revealed distinct patterns of distribution of mRNA (Fig. 1, Table 1) and immunoreactivity (Fig. 2, Table 2) for eight mGluRs in CNS of mammals, especially in rat and mouse. Although only a little difference of functions among splice variants of mGluRs has been reported, there are clear differences of regional and cellular distribution between different splice variants of mGluR1, mGluR7, and mGluR8 (Berthele et al., 1998; Corti et al., 1998; Ferraguti et al., 1998; Kinoshita et al., 1998). Distribution of mRNA and immunoreactivity for mGluR1, mGluR3, mGluR5 and mGluR7 is extensive throughout the brain, whereas that for mGluR2,

OT

7

'. ,i {~. . . . ;7::7{>~~" .; - ""

\ Rt Fig. 1. Distinct distribution of mRNAs for metabotropic glutamate receptor subtypes in the adult rat brain. Parasagittal sections through the brains were hybridized with antisense riboprobes for mGluR1, mGluR2, mGluR3, mGluR4, mGluR5, and mGluR7 as described (Abe et al., 1992; Shigemoto et al., 1992; Ohishi et al., 1993a,b, 1995a,b). AOB, accessory olfactory bulb; Cb, cerebellum; Cx, neocortex; DG, dentate gyms; Hi, hippocampus; MOB, main olfactory bulb; OT, olfactory tubercle; Rt, reticular thalamic nucleus; St, neostriatum; Th, thalamus.

65

Ch. III

R. Shigemoto and N. Mizuno

T A B L E 1. Distribution of mGluR mRNAs in the adult rat CNS Relative grain densities on neuronal cell bodies mGluR 1

mGluR2

mGluR3

mGluR4

mGluR5

mGluR7

Mitral cells

4-4-4-

.

Tufted cells Internal granule cells

4-4-44-

- ~44-

-

+4-~+4-4-

4+

4-++

Periglomerular cells

-

-

-

4-4-

4-

+

4-4-4-

4-4-4-

-

-

-

4-++

44-(s)

4-4+'-,4-++

4-~4-4-

4-44-44-

4-44-4-44-4-

+ ++

Olfactory system Main olfactory bulb .

.

+++

.

Accessory olfactory bulb Mitral cells Granule cells Periglomerular cells Anterior olfactory nucleus Olfactory tubercle Pyramidal cells Islands of Calleja

+

-

- ~+

4-+

++

++

-

-

+

-

++ +

Nucleus of the lateral olfactory tract

4-

4-

- "-'4-

-

4-4-4-

+++

Bed nucleus of the accessory

4-

4-4-

4-

-

4-

+++

4-~4-4-4-

4-~4-4-4-

4-~4-4-4-

+

4-~4-4-4-

4-4-

Piriform cortex

+~+4-+

+

+

4-

4-4-~4-4-4-

4-4-

Cingulate cortex Retrosplenial cortex

4-~4-4-44-~4-4-4-

4-~4-4+"-4-+

4-~4-44-~4-+

Entorhinal cortex Subiculum Presubiculum

4-'--+++ +~++ +'--++

+~+++ +~++ +

+'~++ - "~+

444-4-~4-4-44-4-

4-~4-444-4-~4-4-44-4-4-

4-44-44-44-4-

4-4-

4-4-

4-4-

Parasubiculum Hippocampus

++(s)

+~+++

+~++

4-4-

4-4-4-

4-4-

CA1 pyramidal cells

- "--+

-

-

4-

4-4-4-

4-4-

CA3 pyramidal cells Hilar cells of the dentate gyrus Dentate granule cells

+++ 4-4-44-+

+4-

4-4-

4-44-44-

4-4-44-4-44-4-

4-44-44-4-

Medial septal nucleus Lateral septal nucleus Triangular septal nucleus Septohippocampal nucleus Nuclei of the diagonal band Bed nucleus of the stria terminalis

4-(s) 4-4-44-44-4-44-44-(s)

+ 4-44- ~4-

+

+(s)

+++

4-4-~4-4-44-

4-4-44-4-44-4-4-

4-~4-44-4-

-

+(s)

++

4-

4-

4-4-

Medial preoptic area

4-

4-

-

4-

4-4-

Lateral preoptic area

4-4-'-'4-4-4-

-

-

4-

4-

olfactory tract Neocortex Limbic cortex

Septal and basal forebrain regions

m

m

4-4-

Amygdala Cortical amygdaloid nucleus

4-

_

_ ~ +

_

+~++

++

Medial amygdaloid nucleus

4-~4-+

+

-

_

+

++

Lateral amygdaloid nucleus

4-

++

++

+

++

++

Basolateral amygdaloid nucleus

4-

++

+~++

+

++

Basomedial amygdaloid nucleus Central amygdaloid nucleus

44-

++

-

_

+

++ ++

_

_

+

+ ~ + +

++

+

+++

++

Basal ganglia Striatum

66

4-4-

+(s)

Metabotropic glutamate receptors

Ch. III

TABLE 1 (continued) Relative grain densities on neuronal cell bodies

Nucleus accumbens Globus pallidus Entopeduncular nucleus Ventral pallidum Claustrum Subthalamic nucleus Substantia nigra compact part reticular part Epithalamus Medial habenular nucleus Lateral habenular nucleus Thalamus Anterodorsal nucleus Anteroventral nucleus dorsomedial part ventrolateral part Anteromedial nucleus Mediodorsal nucleus Ventrolateral nucleus Ventromedial nucleus Ventrobasal nuclear complex Gelatinosus nucleus Laterodorsal nucleus Lateroposterior nucleus Paratenial nucleus Paraventricular nuclei Interanteromedial nucleus Intermediodorsal nucleus Rhomboid nucleus Reuniens nucleus Centrolateral nucleus Paracentral nucleus Central medial nucleus Parafascicular nucleus Posterior nuclear group Medial geniculate nucleus Lateral geniculate nucleus Reticular nucleus

mGluR1

mGluR2

mGluR3

mGluR4

mGluR5

mGluR7

+ 4-4-44-4-44-4-44.

+(s) +(s)

+ +

+ -

+++ +~+++

++ + + ++ ++ ++

4.

-

4,

-

4.

+(s)

+

-

+~++(s)

+(s)

-

-

+++

+

-

++

+

4-4-44-4-4-

4"

4"

m

m

__

4.(s)

_

_

_

+

+

4-4-

++

-

++

4,4.4, 4,--~4,4.(s) 4-4-44-4-4+++ 4,4, 4-4-44-+44-4+++ 4-4-4+~+++ 4,4. 4,4. 4-44-44, 4, 4.~4.4. 4-4-44-4-44-4-44-4-

++ +4-

-

"~4"4"

4"

4-44-4+4+44-4++ 4-+

4-44-44" _

-

Zona incerta

4.

Hypothalamus Supraoptic nucleus Paraventricular nucleus Lateral hypothalamic area Suprachiasmatic nucleus Arcuate nucleus Ventromedial nucleus Dorsomedial nucleus Medial mammillary nucleus Lateral mammillary nucleus

4,~4,4,4, +'~++ 4,~4,4,4,(s) 44. 4. 4, 4-44-4-

-

P

++ ++ ++ ++ ++ ++ ++ ++ ++ ++ ++ + ++ ++ ++ ++ ++ ++ ++ ++ ++ ++ +~++

+ ++ +,~++ + + +

+

4.,--.~4.4.

4-4-4-

-

4-+

-

4,

-

4,

++

+(s) + ++ ++ + + 4. -44, 4-4-+ 4, 4,~4.4. +,-~++ + 44.,-~4.4. 4. 4.

4. + 4-4-

+(s) 4-4-

++~+++

+,-~++

++ ++ ++ ++ ++ ++ ++ ++ ++ ++ ++ ++ ++ ++ ++ ++ ++ ++ ++ ++ ++ ++ +44.+ 4-4-

4-44--44-44-44-44-44-44-4-

+

67

Ch. III

TABLE

R. Shigemoto and N. Mizuno

1 (continued) Relative grain densities on n e u r o n a l cell bodies mGluR 1

mGluR2

mGluR3

mGluR4

mGluR5

mGluR7

_

-

_

+

++

-

+

++

+

+(s)

-

+~++

+~++

Midbrain Supramammillary nucleus

++

-

Ventral tegmental area

+(s)

-

Red nucleus

++,~+++

-

-

Interpeduncular nucleus

4.4.~4.4,4,

-

4-4-

4,

Superior colliculus

4-4-

-

4.~+4,

4.

4."~4.4.

4,4,

Inferior colliculus

+(s)

-

++(s)

-

+~++

++ 4-4-

-~+

~+

Oculomotor nucleus

++

-

-

4,

-

Trochlear nucleus

4-+

-

-

+

-

+

Periaqueductal gray Raphe nuclei

+~++ +

-

+(s) _ ~+

+ _

+ _

++ ++

Parabigeminal nucleus

4-4-

-

-

-

+~+4,

4-+

-

4.

-

4-4-

-

4-4-

4,

-

4,

-

4-4-

-

+--~++

++

-

+~++

++

-

4.

4,4,4,

4.

-

4-

+

+

++

~+

Pons and Medulla oblongata Pontine nuclei Pontine reticulotegmental nucleus Parabrachial nuclei

+

-

--~+

-

Dorsal tegmental nucleus

4,

Locus coeruleus

.

Abducens nucleus

4,

-

-

Pontine reticular formation

+

+

-

Mesencephalic trigeminal nucleus

-

-

-

+

-

+

Trigeminal motor nucleus

+(s)

-

-

+

-

+

.

,~+

-

.

.

4. ~+

Principal sensory trigeminal nucleus

+

-

+(s)

+

+

+

Spinal sensory trigeminal nuclei

+

-

+(s)

4,

+~++

+

+(s)

++

Superior olivary complex

+~++

-

Trapezoid body nucleus

4-+

4,

"-~+

++ .

-

Facial nucleus

+(s)

-

-

dorsal nucleus

+++

+(s)

+(s)

+

-

++

ventral nucleus

++

-

+(s)

+

-

++ +~++

.

.

. +

+ -

++ +

Cochlear nuclei

Vestibular nuclei

4-+

+(s)

-

Ambiguus nucleus

-

-

-

N u c l e u s o f t h e s o l i t a r y tr a c t

+(s)

-

-

~+ -~+

-

+

+

4.

+

+

+'~++

++

Dorsal motor vagus nucleus

4.

-

+

-

-

4-

Inferior olivary nuclei

+4-

4,

4,

+

4.'~++

+

Prepositus hypoglossal nucleus

4.

-

-

+

+

Hypoglossal nucleus

++

-

-

4.

-

+

Medullary reticular formation

+

+

-

~+

4,

+

++

Lateral reticular nucleus

-

4-

-

~+

+

-

+

External cuneate nucleus

+

4,

-

~4.

4,

4,

4-4-

Cuneate nucleus

+

-

"~4,

-

~+

+

++

+

Gracile nucleus

+

-

~+

-

"~4.

+

++

+

~+

-

"~+

Cerebellum Cerebellar cortex Purkinje cells

++++

.

granule cells

+

-

-

4-++

-

-

Golgi cells

4-4-

4-4-4-4-

4-4-

-

+4-

4,

stellate cells

++

-

4,

+

-

4,

4-4-

-

4-

+

4,~4,+

4,

Cerebellar nuclei

68

.

.

.

++

Metabotropic glutamate receptors

Ch. III

TABLE 1 (continued) Relative grain densities on neuronal cell bodies

Spinal cord Dorsal horn Ventral horn motor neurons

mGluR 1

mGluR2

mGluR3

mGluR4

mGluR5

mGluR7

+

+(s)

+(s)

+

++

++

++

-

-

++"~+++

Relative grain densities: + + + + = very high; + + + = high; + + = moderate; + -- low; - = background level. The brain regions were demarcated according to Paxinos and Watson (1986). (s) indicates that labeled cells are scattered in each region. Data are adopted and modified from Shigemoto et al. (1992, and unpublished), and Ohishi et al. (1993a,b, 1995a).

sN /

i! j'

Th

GP

s}

./

Vp Fig. 2. Distinct distribution of immunoreactivity for metabotropic glutamate receptor subtypes in the adult rat brain. Parasagittal sections through the brains were reacted with antibodies specific for mGluRla, mGluR1 (all splice variants), mGluR2/3, mGluR4a, mGluR5, mGluR7a, mGluR7b, and mGluR8a as described (Shigemoto et al., 1993, 1997; Ohishi et al., 1994, 1998; Kinoshita et al., 1996a,b, 1998). Acb, nucleus accumbens; AOB, accessory olfactory bulb; Cb, cerebellum; Cx, neocortex; DG, dentate gyms; GP, globus pallidus; Hi, hippocampus; IC, inferior colliculus; LS, lateral septum; MOB, main olfactory bulb; OT, olfactory tubercle; Pir, piriform cortex; Rt, reticular thalamic nucleus; SC, superior colliculus; SN, substantia nigra; SpV, spinal trigeminal nucleus, caudal part; St, neostriatum; Th, thalamus; VP, ventral pallidum.

69

TABLE 2. Distribution of mGluR-like immunoreactivities in CNS Density of immunoreactivities in neuropil mGluR 1

mGluR2

mGluR3

mGluR5

mGluR7a

Olfactory system Main olfactory bulb Glomerular layer External plexiform layer Internal plexiform layer Accessory olfactory bulb Glomerular layer External plexiform and mitral cell layer Granule cell layer Anterior olfactory nucleus Olfactory tubercle Islands of Calleja complex Nucleus of the lateral olfactory tract Bed nucleus of the accessory olfactory tract

++++ ++ +

++ ++ ++

+/+/+/-

+ +++ ++

++ + ++

+++ ++ + + + ++++ + ++

+ +++ +++ - ~++ ++ + ++ +++

+/++ ++ ++~+++ +++ +++ +++ +

+++ ++ +++ ++~+++ ++++ + +§ +

+ +~+++ ++ +~++++ ++++ +++ + +

Neocortex Layer Layer Layer Layer Layer Layer

++ ++ + + + +

+++ +++ +++ +++ ++ ++

+++ +++ +++ +++ +++ +++

+++ +++ ++ ++ ++ +

+++ ++ ++ + ++ +

+ + + +~+++ + +

+ +++ +++ +++ ++ +++

+++ +++ +++ +++ + +++

++ ++ ++ ++~+++ +++ ++++

+++~++++ ++ + ++~++++ + ++

+/+/+/-(c, +++)

+++ + +

++ ++ ++

+++ ++++ ++++

++ +++ +++

I II III IV V VI

Limbic cortex Piriform cortex Cingulate cortex Retrosplenial cortex Entorhinal cortex Tenia tecta Subiculum Hippocampus CA1, stratum lacunosum-moleculare CA1, stratum radiatum CA1, stratum oriens

mGluR7b

m

m

m

+ +§247

~z

t,,~~

T A B L E 2 (continued) Density of immunoreactivities in neuropil

CA3, stratum lacunosum-moleculare CA3, stratum radiatum CA3, stratum lucidum CA3, stratum oriens Hilus of the dentate gyrus Molecular layer of the dentate gyrus Septal and basal forebrain regions Medial septal nucleus Lateral septal nucleus Triangular septal nucleus Nucleus of the diagonal band Septofimbrial nucleus Bed nucleus of the stria terminalis Substantia innominata Medial preoptic area Lateral preoptic area Amygdala Periamygdaloid cortex Cortical amygdaloid nucleus Medial amygdaloid nucleus Lateral amygdaloid nucleus Basolateral amygdaloid nucleus Central amygdaloid nucleus Basal ganglia Striatum Nucleus accumbens Globus pallidus Entopeduncular nucleus Ventral pallidum Claustrum Subthalamic nucleus ...j

mGluR 1

mGluR2

mGluR3

mGluR5

mGluR7a

mGluR7b

++

+ + +

+ + +

+ + +

+ + +

-

+ + ~ + + +

-

++

+++ ++ ++

-

++ +++

++~+++ + +++ + +++

++ + ++ +++ +++

-

+

++ + ++ + +++

+ +~+++ + +

+ ++ +

+ +++ + -

-~+ ++++ +++ +

_ +~++ ++ -

++

-

+

++

+

-

+(c,+++)

+(c,+++) +(c,+++) +(c,+++)

+ + +

++ + + +

+~++ + + +

+~++ ++ + +

-~++ +++ -

+ + + + +~++ +

+++ ++ ++ ++ +++ ++

+ + + +++ +++ ++

++ +~++ + + + - ~+

++~++++ + + + ++ +~++

+++ +

+++ -

_ _

-

m

++(c,+++) ++ ++

+ + +

+ + +

+ ++ +

++~+++ + -

+§ + +++

~,,~.

"---I

TABLE

2 (continued) Density of immunoreactivities mGluR1

Substantia

mGluR2

in n e u r o p i l mGluR3

mGluR5

mGluR7a

mGluR7b

nigra

pars compacta

+++

+

.

pars reticulata

+

+

+~++

+

++

++

+

§

§

§

+

-

Peripeduncular

nucleus

.

.

.

Epithalamus habenular

nucleus

-

§247

-

-

++§

Lateral habenular

nucleus

§

§

§

§

§

-

Medial

~+

[-]

-

Thalamus Anterodorsal

++

++

+

+/-

+

-

Anteroventral

nucleus nucleus

++

+++

+

+

+ +

-

Anteromedial

nucleus

++

++

+

+

+

-

Mediodorsal

nucleus

+++

+

+

+

-

-

Ventrolateral

nucleus

+++

§

§

+

-

-

§

+

§

§

-

-

+++

+

+

+

+

+

+++

-

+

+

-

+

+++

++

+

+~++

+

-

+ + +

+ +

+

+

+ ~ + +

-

++

+

+

+

-

-

+ ~ + +

+ + +

+

+

+

-

++

++

+

+

-

-

++

++

+

+

-

-

++

++

+

+

-

-

++

++

+

++

-

-

§

§

§

§

-

-

+

+

+

+

-

-

Ventromedial

nucleus

Ventrobasal

nuclear

Gelatinosus

nucleus

Laterodorsal

nucleus

Lateroposterior Paratenial

complex

nucleus

nucleus

Paraventricular

nuclei

Interanteromedial Intermediodorsal Rhomboid

nuclei nuclei

nucleus

Reuniens

nucleus

Centrolateral Paracentral

nucleus nucleus

Centromedial

nucleus

+

§247

§

§

-

-

Parafascicular

nucleus

§247

-

§

§

-

-

+§

-

§

§

-

-

Posterior Medial

nuclear

geniculate

group nucleus

Dorsal lateral geniculate Intergeniculate

leaflet

nucleus

-

§

-

+~++

-

-

++§

-

+

~+

§

++

-

+

-

+

§

-

-

~,~~

T A B L E 2 (continued) Density of i m m u n o r e a c t i v i t i e s in neuropil

Ventral lateral g e n i c u l a t e nucleus R eticular n u c l e u s

mGluR1

mGluR2

+++

+

+/-

+~+++

Z o n a incerta

+~++

+

Hypothalamus Supraoptic n u c l e u s

+(c,

mGluR5

mGluR7a

+§ +/-

+~++

__

+

,-,., §

mGluR7b ~o

+

o~

-

+

-

+

-

+

-

+

-

++(c, +++)

+

+

+

+

+

Paraventricular nucleus

+/-(c,

Lateral h y p o t h a l a m i c area

+++)

mGluR3

+++)

-

++

-

+

+ / -

-

_

+ / _

-

+

-

+

-

+

-

+

-

+

-

+

+

+

++

+

-

+ + +

+ -

+ + +

+ + +~++

+ + +

-

++

++

+

+~++

- ~++

-

Lateral m a m m i l l a r y nucleus

+

-

+

+/-

+

-

S u p r a m a m m i l l a r y nucleus

++

-

+

+

++

-

+~++ +++

-

+ +

+ ++

+ +

Ventral t e g m e n t a l area

+

+

- ~+

-

Red nucleus I n t e r p e d u n c u l a r nucleus

++

.

Rostral subdivision

+

++

+

+

+

Lateral subdivision

+

++

+

+

+

D orsolateral subdivision

++

-

+

+

-

D o r s o m e d i a l subdivision

+

-

+

+

-

C a u d a l subdivision

+

++

+

+

-

S u p r a c h i a s m a t i c nucleus Periventricular nucleus Arcuate nucleus Ventrome~tial n u c l e u s Dorsomedial nucleus Posterior h y p o t h a l a m i c nucleus P r e m a m m i l l a r y nucleus Medial m a m m i l l a r y nucleus

Midbrain Pretectum Pretectal olivary nucleus

.

.

.

S u p e r i o r colliculus Superficial layer

+++

+

+

++

++++

I n t e r m e d i a t e layer

+

+

+

+

+

D e e p layer

+

+

+

+

-

4~

TABLE

2 (continued) Density of immunoreactivities

in n e u r o p i l

mGluR 1

mGluR2

Inferior colliculus

+~++

-

Oculomotor

+

.

.

.

.

Trochlear

nucleus

Cuneiform Median

+

nucleus

Periaqueductal

gray

+(c, +

nucleus

raphe nucleus

+

nuclei

Pontine

reticulotegmental

tegmental

tegmental

Dorsal tegmental

nucleus

nucleus nucleus

Ventral tegmental Locus

nucleus

nucleus

Pedunculopontine

Laterodorsal

nucleus

tegmental

nucleus

coeruleus

Pontine

reticular formation

Trigeminal Principal

.

.

.

-

-

. .

++

+

-

+

-

+

~+

-

+

-

+

-

+/-

+

-

+++

-

-

+"~++

+

-

-

++

+

-

+

-

+++)

~+

+ +

oblongata

Pontine

Anterior

+~++ .

mGluR7b

+

Parabigeminal

Parabrachial

+

mGluR7 a

-

+(c,

nucleus

mGluR5

-

Dorsal raphe nucleus

Pons and Medulla

+++)

mGluR3

motor nucleus sensory

trigeminal

Spinal trigeminal

nucleus

~+

+

.

++ .

.

.

.

+

-

+

+

+

.

++

+++

-

+

+

++

-

+

+++

-

-

++

+++

-

+

++

-

+

-

+

+

-

-

+

-

+/-

+

++++

+

+

-

-

-

+

-

+

.

++

+

+~++ .

.

.

.

+ .

.

++

. +

nucleus

Oral subnucleus

+

++

+

+

-

-

Interpolar

+

++

+

+

-

-

superficial laminae

++

++

+

+++

+++

+

deeper laminae

+

+

+

+

+

-

Caudal

Superior Trapezoid

subnucleus

subnucleus

olivary complex

+

+

+

-

-

-

body nucleus

+

+

-

+

-

-

Facial nucleus Dorsal cochlear Ventral cochlear

nucleus nucleus

+

.

+++

++~-,+++

. +

.

. -

-

-

++

++

+

-

-

-

t,,,,

. t-.I

TABLE 2 (continued) D e n s i t y of i m m u n o r e a c t i v i t i e s in n e u r o p i l t...,

mGluR1

mGluR2

mGluR3

mGluR5

mGluR7a

mGluR7b t...,.

Vestibular nucleus lateral n u c l e u s

+

+

+/-

+/-

-

+, [++]

medial nucleus

++

+

+

+

-

-

superior nucleus

++

-

+/-

+

-

-

spinal n u c l e u s

++

-

+

+/-

-

-

+ +

+

+ +

+ - ~++

+ ++

-

++

+

Ambiguus nucleus N u c l e u s of the solitary tract Dorsal motor vagus nucleus

+

.

I n f e r i o r olivary n u c l e i

+++

++

.

.

Hypoglossal nucleus Medullary reticular formation

+ +

. .

-

-

Lateral reticular nucleus

+

++

+

-

+

-

External cuneate nucleus

+

++

+

+

-

-

Cuneate nucleus

+

-

+

+

-

-

Gracile nucleus

+

-

+

+

-

-

Area postrema

++

.

++ . .

.

+~++

. .

. .

. +

.

.

.

.

.

Cerebellum Cerebellar cortex molecular layer P u r k i n j e cell l a y e r g r a n u l e cell l a y e r Cerebellar nuclei

-

+

-

+ + +

+

+

+

+

-

_

_

+

+(c, + + + ) ++

+++ +

+/+/-

+/+~++

++, [+++]

++ +

++ +

+ +/-

++ +

-

+/-

Spinal cord Dorsal horn Intermediate zone Ventral h o r n I n t e n s i t y of i m m u n o r e a c t i v i t y : + + + +

+ = m o s t intense; + + +

= intense; + +

= m o d e r a t e ; + -- w e a k ; + / -

+++

+

+

-- v e r y w e a k ; -

= negative. (c, + + + )

indicates intensely

l a b e l e d cells s c a t t e r e d in e a c h region. D a t a are o b t a i n e d f r o m the rat e x c e p t those for m G l u R 3 , w h i c h w e r e o b t a i n e d f r o m m G l u R 2 - d e f i c i e n t m i c e u s i n g an a n t i b o d y to m G l u R 3 w i t h s o m e c r o s s - r e a c t i v i t y to m G l u R 2 (Y. T a m a r u et al., u n p u b l i s h e d ) . [ ] in m G l u R 7 a and m G l u R 7 b c o l u m n s i n d i c a t e s data in the m o u s e , w h e n t h e y are d i f f e r e n t f r o m those in the rat. D a t a are a d o p t e d a n d m o d i f i e d f r o m S h i g e m o t o et al. (1993, and u n p u b l i s h e d ) , O h i s h i et al. (1998), Y. T a m a r u et al. ( u n p u b l i s h e d ) , a n d K i n o s h i t a et al. (1998). ----..I

o~

Ch. III

R. Shigemoto and N. Mizuno

mGluR4 and mGluR8 is found relatively restricted to specific brain regions. In the adult rat, most of the mRNA signals for mGluRs are observed in neuronal cells except those for mGluR3, which is extensively expressed in glial cells throughout brain regions (Ohishi et al., 1993b, 1994; Tanabe et al., 1993; Testa et al., 1994; Makoff et al., 1996b; Petralia et al., 1996a; Mineff and Valtschanoff, 1999). However, expression of mGluR5 in some astrocytes has been found in the hypothalamus of adult rats (Van den Pol et al., 1995), and in the thalamus and hippocampus of young rats (Liu et al., 1998; Schools and Kimelberg, 1999). Expression of group I and group II mGluRs in other types of glial cells was also reported in ependymal cells (mGluRla: Tang and Sim, 1997), interstitial glial cells of the pineal gland (mGluR2/3 and mGluR5: Pabst and Redecker, 1999), and pinealocytes (mGluR5: Yatsushiro et al., 1999). Expression of mGluR6 has been found only in the retina but not in the brain or spinal cord (Nakajima et al., 1993; Nomura et al., 1994; Schools and Kimelberg, 1999). Regional distribution of mRNA and immunoreactivity for group I mGluRs correspond very well reflecting that these mGluR proteins are mostly localized in somatodendritic domains of neurons, near the site of protein synthesis. On the other hand, group II mGluRs are observed not only in somatodendritic domains but also in axonal domains, and group III mGluRs, except for mGluR6, are present mainly in axon terminals as described in detail in Section 3. These situations make regional distribution of immunoreactivity for group II and group III mGluRs sometimes quite different from that of mRNAs. For example, the most intense immunoreactivity for mGluR2 is observed in the neuropil of the stratum lacunosum moleculare of the hippocampal CA1 area (Ohishi et al., 1998), whereas no expression of mGluR2 mRNA was detected in the CA1 area (Ohishi et al., 1993a). Similarly, immunoreactivity for mGluR4a is abundant in the globus pallidus (Bradley et al., 1999), but no mRNA for mGluR4a was detected there (Ohishi et al., 1995a). In both cases, lesions generated in the entorhinal cortex and neostriatum, which send massive projection fibers to the CA1 area and globus pallidus, respectively, markedly reduced immunoreactivity for the respective mGluRs (Shigemoto et al., 1997; Bradley et al., 1999), indicating transport of the receptor proteins to presynaptic elements. 2.2. DISTRIBUTION OF mRNA AND IMMUNOREACTIVITY FOR GROUP I METABOTROPIC GLUTAMATE RECEPTORS 2.2.1. mGluR1 mRNA

Distribution of mGluR1 mRNA in the CNS was investigated in the rat by in situ hybridization histochemistry (Masu et al., 1991; Shigemoto et al., 1992; Fotuhi et al., 1994; Testa et al., 1994; Kerner et al., 1997). According to a systematic study (Shigemoto et al., 1992), mGluR1 mRNA was distributed widely throughout the CNS (Table 1): most intense expression was seen in Purkinje cells of the cerebellar cortex, mitral and tufted cells of the olfactory bulb, granule cells of the dentate gyrus, neurons in the hilus, pyramidal neurons of CA3, as well as neurons in the lateral septum, globus pallidus, entopeduncular nucleus, ventral pallidum, magnocellular preoptic nucleus, substantia nigra pars compacta and pars reticulata, and dorsal cochlear nucleus. Neurons showing moderate expression were seen in high density in the superficial layers of the cingulate, retrosplenial and entorhinal cortices, dentate gyrus, islands of Calleja, mammillary nuclei, red nucleus, and superior colliculus. The expression was detected in most of the thalamic neurons, but not in the thalamic reticular nucleus. In the developing rat brain, the level of mGluR1 mRNA gradually increased during early postnatal days according to the maturation of neuronal elements. However, in the lumber cord of the rat, 76

Metabotropic glutamate receptors

Ch. III

it was reported that the expression of mGluR1 mRNA was generally decreased from postnatal day 1 to postnatal day 21 (Berthele et al., 1999). In the rat retina, expression of mGluR1 mRNA was observed with moderate intensity in the large majority of neurons in the ganglion cell layer, suggesting that both ganglion cells and a subset of amacrine cells expressed mGluR1 mRNA; moderate expression was also seen in some putative amacrine cells with cell bodies in the inner third of the inner nuclear layer (Hartveit et al., 1995). Differential expression of mRNAs for mGluR1 splice variants was observed in the rat (Pin et al., 1992; Berthele et al., 1998) and human (Berthele et al., 1998). In the rat, the mGluRld mRNA was expressed widely and the mGluRla and mGluRlb mRNAs were expressed in almost complementary patterns. On the other hand, formation of mGluRlc splice variants appeared to be a rare event (also see Pin et al., 1992). Strong expression of the mGluRla mRNA was seen in Purkinje cells, the mitral and tufted cells, hippocampal interneurons, thalamic neurons, and neurons in the substantia nigra, and moderately expressed in the superior and inferior colliculi and cerebellar granule cells. The mGluRlb mRNA was expressed strongly in Purkinje cells, hippocampal pyramidal neurons, granule cells of the dentate gyrus, and lateral septum, and also was expressed moderately in neurons in the striatum and superficial layers of the cerebral cortex, as well as in granule cells of the cerebellar cortex. The mGluRld mRNA was expressed in all regions where the mGluRla and mGluRlb mRNAs were detected; it was strongly expressed in Purkinje cells, mitral and tufted cells, pyramidal neurons and interneurons in the hippocampus, and neurons in the thalamus and substantia nigra, and also was expressed moderately in the lateral septum, cerebral cortex, striatum, and superior and inferior colliculi. In human, mGluR1 splice variant expression in the cerebellum was also found to match that observed in the rat.

2.2.2. mGluR1 immunoreactivity Distribution of immunoreactivity for mGluR1 (for all mGluR1 splice variants) and mGluRla was observed extensively in the rat brain regions (Martin et al., 1992; Baude et al., 1993; Fotuhi et al., 1993; Petralia et al., 1997). In the hippocampus of the rat, immunoreactivity for mGluR1 was strong in dendritic fields of the dentate gyrus and CA3 area. Intensely immunoreactive intemeurons are also scattered in the hilus and CA areas being most densely distributed in the border region between the CA1 stratum oriens and alveus. On the other hand, mGluRla immunoreactivity was found only in the interneurons indicating that the mGluR1 immunoreactivity in the dendritic fields of dentate granule cells and CA3 pyramidal neurons is ascribable to expression of mGluRlb, mGluRlc and/or mGluRld (Shigemoto et al., 1997). Much stronger immunoreactivity for mGluR1 than that for mGluRla is also apparent in the lateral septum, islands of Calleja, supraoptic nucleus, paraventricular nucleus, and some scattered neuronal cell bodies in the preoptic areas, lateral hypothalamus, and central amygdaloid nucleus. It is reported that some of these regions have strong immunoreactivity for mGluRlb (Ferraguti et al., 1998; Mateos et al., 1998). Distribution of immunoreactivity for mGluR1, mGluRla and mGluRlb was further studied in the forebrain of the rat and mouse (Ferraguti et al., 1998), piriform cortex and olfactory tubercle of the rat (Wada et al., 1998), hippocampus of the rat (Hampson et al., 1994), basal ganglia of the rat (Tallaksen-Greene et al., 1998; Testa et al., 1998), lateral geniculate nucleus of the cat (Godwin et al., 1996), hypothalamus of the rat (Van den Pol, 1994; Van den Pol et al., 1994; Mateos et al., 1998), cerebellum of the rat (G6rcs et al., 1993; Grandes et al., 1994; Hampson et al., 1994; Jaarsma et al., 1998), dorsal cochlear nucleus of the rat (Petralia et al., 77

Ch. III

R. Shigemoto and N. Mizuno

1996b; Jaarsma et al., 1998), and autonomic cell groups of the medulla oblongata of the rat (Hay et al., 1999). In the human cerebral cortex, the presence of mGluRla immunoreactivity was reported in a small number of non-pyramidal cells, but not in pyramidal neurons (Ong et al., 1998). Comparison of the results obtained from in situ hybridization histochemistry with those from immunohistochemistry indicated that the mGluR1 was expressed in the somatodendritic domain of neurons but not in axons in the brain and spinal cord. In fact, expression of mGluR1 in postsynaptic neuronal elements was confirmed electron-microscopically in the cerebral cortex (Ong et al., 1998), hippocampus (Baude et al., 1993; Lujfin et al., 1996, 1997; Hanson and Smith, 1999), striatum (Hanson and Smith, 1999), thalamus (Martin et al., 1992; Godwin et al., 1996; Liu et al., 1998), hypothalamus (Van den Pol, 1994), cerebellar cortex (Martin et al., 1992; Baude et al., 1993; G6rcs et al., 1993; Nusser et al., 1994; Lujfin et al., 1996, 1997; Jaarsma et al., 1998), and dorsal cochlear nucleus (Petralia et al., 1996b; Jaarsma et al., 1998). In the rat retina, mGluRla immunoreactivity light-microscopically was observed mostly in the inner plexiform layer (Peng et al., 1995) or in the outer and inner plexiform layers (Koulen et al., 1997). Electron-microscopically, mGluRla immunoreactivity in the outer plexiform layer was seen in rod bipolar cell dendrites postsynaptic at ribbon synapses of rod photoreceptor cells, while mGluRla immunoreactivity in the inner plexiform layer was observed in thin amacrine cell processes postsynaptic to OFF-cone bipolar cell terminals, ON-cone bipolar cell terminals, and rod bipolar cell terminals (Koulen et al., 1997). In the cat retina, mGluRla immunoreactivity was observed in the rod spherules in the outer plexiform layer, as well as in amacrine and ganglion cell somata with processes ramifying throughout the inner plexiform layer (Cai and Pourcho, 1999). The developmental changes of mGluRla immunoreactivity was also reported in the visual cortex of the cat (Reid et al., 1995), in the thalamus of the mouse (Liu et al., 1998), in the trigeminal nuclei, ventral posterior thalamic nucleus and barrel area of the somatosensory cortex of the mouse (Mufioz et al., 1999), and in the retina of the rat (Koulen et al., 1997). 2.2.3. mGluR5 mRNA

A wide distribution of the mRNA for mGluR5 throughout the CNS was shown in the rat by in situ hybridization histochemistry (Fig. 1, Table 1; Abe et al., 1992). Intense expression was seen mainly in the telencephalic regions, including the cerebral cortex, hippocampus, subiculum, internal granular layer of the olfactory bulb, anterior olfactory nucleus, pyramidal cell layer of the olfactory tubercle, striatum, accumbens nucleus, and lateral septal nucleus. Strong expression was also seen in the anterior thalamic nuclei, shell regions of the inferior colliculus, and caudal subnucleus of the spinal trigeminal nucleus. In these regions, mGluR5 mRNA was expressed intensely in most neuronal cell bodies. In the hippocampus, neuronal cell bodies showing intense expression of mGluR5 mRNA were distributed throughout the CA pyramidal cells and granule cells in the dentate gyms. In the cerebellar cortex, only a small population (10%) of Golgi cells expressed mGluR5 mRNA; no expression was detected in Purkinje cells or granule cells, although weak expression was seen in these cells in the 6-day-old rat. Expression of mGluR5 mRNA was also studied in the neocortex (Kerner et al., 1997), entorhinal cortex (Fotuhi et al., 1994), hippocampus (Fotuhi et al., 1994; Kerner et al., 1997), and striatum of the rat (Testa et al., 1994, 1995). It was reported that mGluR5 mRNA was expressed intensely in neocortical and hippocampal neurons with immunoreactivity for glutamic acid decarboxylase (Kerner et al., 1997), as well as in striatal projection neurons showing substance P immunoreactivity and enkephalin immunoreactivity. 78

Metabotropic glutamate receptors

Ch. III

In the lumber cord of the rat, it was reported that the expression of mGluR5 mRNA was marked at birth, especially in the superficial dorsal horn, but that the expression levels decreased with age (Berthele et al., 1999). In the rat retina, mGluR5 mRNA was expressed in the outer one third to one half of the inner nuclear layer; it was presumed that mGluR5 mRNA was expressed in the horizontal cells and also in some bipolar cells (Hartveit et al., 1995). Distribution of mRNA specific for mGluR5b also was examined by in situ hybridization (Joly et al., 1995): intense expression was seen in the hippocampus, striatum, lateral septal nucleus, and cerebral cortex. This distribution pattern of mGluR5b mRNA corresponded exactly to that of mGluR5 mRNA (Abe et al., 1992). However, it appeared that mRNA for mGluR5a was most abundant in the young rat, while that for mGluR5b was predominant in the adult rat (Joly et al., 1995; Romano et al., 1996).

2.2.4. mGluR5 immunoreactivity Distribution of mGluR5 immunoreactivity was examined systematically in the rat brain (Shigemoto et al., 1993; Romano et al., 1995). According to the study using an antibody against a fusion protein containing a C-terminal sequence of rat mGluR5 (Table 2; Shigemoto et al., 1993), most intense immunoreactivity was observed in the accessory olfactory bulb, olfactory tubercle, lateral septum, striatum, accumbens nucleus and the CA1 area of the hippocampus. Intense immunoreactivity was also seen in the main olfactory bulb, anterior olfactory nuclei, cerebral cortex, CA3 and dentate gyrus, shell regions of the inferior colliculus, superficial layers of the superior colliculus, and caudal subnucleus of the spinal trigeminal nucleus. Although the neuropil of the striatum showed intense immunoreactivity, immunoreactivity in the cytoplasm of striatal neurons was rather weak. Similarly, in the hippocampus, dendritic fields showed intense immunoreactivity, whereas the pyramidal and granule cell layers were devoid of immunoreactivity (also see Shigemoto et al., 1997). In the cerebellum, 10% of the Golgi cells showed mGluR5 immunoreactivity (Neki et al., 1996b), in accordance with the findings that only a small population of Golgi cells expressed mGluR5 (Abe et al., 1992). Thus, the results of the immunohistochemical study of mGluR5 corresponded well with those of in situ hybridization histochemistry for mGluR5 mRNA (Tables 1 and 2). Distribution of mGluR5 immunoreactivity was further examined in the olfactory tubercle and piriform cortex (Wada et al., 1998), hippocampus (Shigemoto et al., 1997), striatum (Tallaksen-Greene et al., 1998), cerebellum (Neki et al., 1996b; N6gyessy et al., 1997), parabrachial and K611iker-Fuse nuclei (Guthmann and Herbert, 1999), dorsal cochlear nucleus (Petralia et al., 1996b), autonomic cell groups of the medulla oblongata (Hay et al., 1999), and spinal dorsal horn of the rat (Jia et al., 1999), as well as in the lateral geniculate nucleus of the cat (Godwin et al., 1996), and thalamus of the developing mouse (Liu et al., 1998). Immunoreactivity for mGluR5a was also observed in the spinal dorsal horn of the rat (Vidny~nszky et al., 1994). Electron-microscopical studies indicated that mGluR5 was mainly localized in somatic and dendritic profiles: expression of mGluR5 immunoreactivity was observed in the postsynaptic elements in the hippocampus (Luj~n et al., 1996, 1997; Hanson and Smith, 1999), basal ganglia (Shigemoto et al., 1993; Hanson and Smith, 1999), thalamus (Godwin et al., 1996; Liu et al., 1998), hypothalamus (Romano et al., 1995; Van den Pol et al., 1995), cerebellar cortex (N6gyessy et al., 1997), dorsal cochlear nucleus (Petralia et al., 1996b), and dorsal horn of the spinal cord (Vidny~nszky et al., 1994; Jia et al., 1999). 79

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In a few studies, mGluR5 immunoreactivity was reported not only in somatodendritic domains of neurons but also in axons (Romano et al., 1995) or vesicle-containing profiles (Jia et al., 1999), as well as in astrocytes (Van den Pol et al., 1995). In the rat retina, mGluR5 immunoreactivity in the outer and inner plexiform layers was further observed electron-microscopically: immunoreactivity in the outer plexiform layer was in dendrites of rod bipolar cells postsynaptic to rod photoreceptor terminals, while that in the inner plexiform layer was seen in amacrine cell processes postsynaptic to OFF-cone bipolar cell terminals, ON-cone bipolar cell terminals, and rod bipolar cell terminals (Koulen et al., 1997). Developmental changes of mGluR5 immunoreactivity was reported in the visual cortex of the cat (Reid et al., 1995), thalamus of the mouse (Liu et al., 1998), hypothalamus of the rat (Van den Pol et al., 1995), trigeminal nuclei, ventral posterior thalamic nucleus and barrel area of the somatosensory cortex of the mouse (Mufioz et al., 1999). According to a Western blotting analysis, there was more mGluR5 protein present in the brain regions in the developing rat than in the adult (Romano et al., 1996; also see Joly et al., 1995). 2.3. DISTRIBUTION OF mRNA AND IMMUNOREACTIVITY FOR GROUP II METABOTROPIC GLUTAMATE RECEPTORS 2.3.1. mGluR2 mRNA

Distribution of the mRNA for mGluR2 in the CNS was examined in the rat by in situ hybridization histochemistry (Tanabe et al., 1992; Ohishi et al., 1993a). The mRNA for mGluR2 was distributed in more limited regions in the CNS than mRNAs for mGluR1, mGluR5, and mGluR3. According to a systematic study (Ohishi et al., 1993a), the most intense expression of mGluR2 mRNA was observed in Golgi cells in the cerebellar cortex. Strong expression was seen in the mitral cells of the accessory olfactory bulb, external part of the anterior olfactory nucleus, some cells in the entorhinal and parasubicular cortices. Moderate expression was observed in the granule cells of the accessory olfactory bulb, some neurons in the anterior olfactory nucleus, some neurons in the neocortex, cingulate, retrosplenial, and subicular cortices, granule cells of the dentate gyrus, triangular septal nucleus, lateral, basolateral and basomedial amygdaloid nuclei, medial mammillary nucleus, some part of the thalamus including the anterior, ventrolateral, midline, intralaminar, and centromedian-parafascicular thalamic nuclei, retinal ganglion cells, and some cells which were scattered in the inner part of the inner nuclear layer of the retina. Expression of mGluR2 mRNA also was observed in the dentate gyrus and inner layer of the entorhinal cortex (Fotuhi et al., 1994) and striatum (Testa et al., 1994) of the rat. In the rat retina, expression of mGluR2 mRNA was reported in some cells in the ganglion cell layer and inner third of the inner nuclear layer; some of the ganglion cells and a subset of amacrine cells were presumed to express mGluR2 mRNA (Hartveit et al., 1995). 2.3.2. mGluR3 mRNA

Expression of mGluR3 mRNA was seen widely throughout the CNS of the rat by in situ hybridization histochemistry (Ohishi et al., 1993b; Tanabe et al., 1993). According to a systematic study (Ohishi et al., 1993b), the expression was marked in the cerebral cortex, granule cell layer of the dentate gyrus, lateral and basolateral amygdaloid nuclei, dorsal endopiriform nucleus, thalamic reticular nucleus, supraoptic nucleus, superficial layers of the superior colliculus, and Golgi cells in the cerebellar cortex; the expression was most prominent 80

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in the thalamic reticular nucleus neurons, and moderate in Golgi cells. Glial cells also appeared to express mGluR3 mRNA in many regions including the corpus callosum and anterior commissure. Expression of mGluR3 mRNA was also reported in the cerebral cortex, striatum, thalamus, and cerebellum of hiJman (Makoff et al., 1996b), as well as in the dentate gyrus and inner layer of the entorhinal cortex (Fotuhi et al., 1994), striatum (Testa et al., 1994) and spinal cord (Boxall et al., 1998) of the rat. It was reported in the basal ganglia of the rat that mGluR3 mRNA was expressed in glia in all basal ganglia structures, and that mGluR mRNA expression in neurons was observed only in the striatum, nucleus accumbens, substantia nigra pars reticulata, and very weakly in the subthalamic nucleus (Testa et al., 1994). In the lumber spinal cord of the rat, the expression levels and the regional distribution of mGluR3 mRNA were reported to be altered with postnatal development. The expression level of mGluR3 mRNA was highest at birth, especially in the superficial dorsal horn, but these levels decreased with age; up to postnatal day 12, the expression was almost exclusively restricted to the grey matter, but with postnatal day 21 a strong additional expression occurred in the white matter (Berthele et al., 1999). No expression of mGluR3 mRNA was detected in the retina (Hartveit et al., 1995).

2.3.3. mGluR2/3 immunoreactivity Distribution of group II mGluRs in the brain and spinal cord was studied by using polyclonal antibodies against C-terminus of rat mGluR2; these antibodies recognized both mGluR2 and mGluR3 (Ohishi et al., 1994; Petralia et al., 1996a). An extensive study of distribution of mGluR2/3 immunoreactivity in the rat brain and spinal cord was reported (Petralia et al., 1996a): light-microscopical distribution of mGluR2/3 immunoreactivity matched the combined distributions of mGluR2 mRNA and mGluR3 mRNA; the most intense immunoreactivity was seen in presumptive necklace olfactory glomeruli neurons of the superficial glomeruli of the accessory olfactory bulb, Golgi cells of the cerebellar cortex, and border region between lamina II and lamina III of the spinal dorsal horn. In the hippocampus, the immunoreactivity was most strong in the neuropil of the CA3 stratum lucidum/pyramidale, CA1 and CA3 stratum lacunosum moleculare, hilus and middle one third of the molecular layer of the dentate gyrus. Electron-microscopy revealed mGluR2/3 immunoreactivity in postsynaptic and presynaptic structures and glial wrappings of synapses in the cerebral cortex, hippocampus, and striatum; in the hippocampus, mGluR2/3 immunoreactivity in the presynaptic structures was concentrated in axon terminals of two populations of presumptive glutamatergic axons: mossy fibers and perforant path (Petralia et al., 1996a). Distribution of mGluR2/3 immunoreactivity in particular regions of the CNS was also examined, in the hippocampus (Shigemoto et al., 1997), basal ganglia (Testa et al., 1998), cerebellar cortex (Ohishi et al., 1994; Neki et al., 1996b; Jaarsma et al., 1998), parabrachial and K611iker-Fuse nuclei (Guthmann and Herbert, 1999), cochlear nuclei (Petralia et al., 1996b; Jaarsma et al., 1998), autonomic cell groups of the medulla oblongata (Hay et al., 1999), and spinal dorsal horn (Jia et al., 1999) of the rat, as well as in the retina of the rat (Koulen et al., 1996) and cat (Cai and Pourcho, 1999). Immunoreactivity for mGluR2/3 was observed electron-microscopically not only in somatodendritic neuronal domains but also in axonal domains (Hayashi et al., 1993; Ohishi et al., 1994; Petralia et al., 1996a,b; Yokoi et al., 1996; Lujfin et al., 1997; Shigemoto et al., 1997; Jaarsma et al., 1998; Liu et al., 1998; Wada et al., 1998; Cai and Pourcho, 1999; Jia et al., 1999; Meguro et al., 1999), particularly at both presynaptic and postsynaptic elements of Golgi cells of the cerebellar cortex (Ohishi et al., 1994; Neki et al., 1996a,b) and in the dendro81

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dendritic synapses of the granule cells in the accessory olfactory bulb (Hayashi et al., 1993), and mainly presynaptically in the hippocampus (Petralia et al., 1996a; Lujfin et al., 1997; Shigemoto et al., 1997; Ohishi et al., 1998). The existence of mGluR 2/3-immunoreactivity was also reported in Bergmann glia in postnatal rat (Meguro et al., 1999) as well as in astrocytes (Petralia et al., 1996a,b; Liu et al., 1998; Mineff and Valtschanoff, 1999). In the rat hippocampus, mGluR2/3 immunoreactivity was strong in terminal zones of the mossy fibers and perforant path, and was most dense in the lacunosum moleculare of the CA1 area; electron-microscopically it was found frequently in small unmyelinated axons, especially in preterminal portions of axons rather than in axon terminals (Yokoi et al., 1996; Shigemoto et al., 1997). In the retina of the rat, mGluR 2/3 immunoreactivity was localized exclusively in the processes of cholinergic amacrine cells, which were postsynaptic to bipolar cell synapses in the inner plexiform layer (Koulen et al., 1996). In the retina of the cat, the immunoreactivity was observed in horizontal cells and amacrine cells; the immunoreactive amacrine processes were postsynaptic to cone bipolar cells and to rod bipolar terminals (Cai and Pourcho, 1999). Developmental changes of mGluR2/3 immunoreactivity were examined in the thalamus of the mouse (Liu et al., 1998), in the trigeminal nuclei, ventral posterior thalamic nucleus and barrel area of the somatosensory cortex of the mouse (Mufioz et al., 1999), in the cerebellar cortex of the rat (Meguro et al., 1999), and in the retina of the rat (Koulen et al., 1996). In the cerebellar cortex, Bergmann glial cells with their radial processes into the molecular layer showed mGluR2/3-immunoreactivity during the early postnatal period (Meguro et al., 1999).

2.3.4. mGluR2 immunoreactivity Distribution of mGluR2 in the brain and spinal cord was examined immunohistochemically in the rat and mouse by using a monoclonal antibody that was raised against an N-terminal sequence of rat mGluR2 (amino acid residues 87-134) (Table 2; Neki et al., 1996a; Ohishi et al., 1998): the distribution pattern of mGluR2-immunoreactive neuronal cell bodies (Ohishi et al., 1998) was in good accordance with that of mGluR2 mRNA (Ohishi et al., 1993a). It was indicated, however, that mGluR2 was located not only in the somato-dendritic domain but also in the axonal domain of neurons; no glial cells showing mGluR2-immunoreactivity were found. The neuropil was intensely immunostained in the accessory olfactory bulb, bed nucleus of the accessory olfactory tract, cerebral neocortex, cingulate cortex, retrosplenial cortex, subicular and entorhinal cortices, stratum lacunosum moleculare of CA1-3, molecular layer of the dentate gyrus, periamygdaloid cortex, basolateral amygdaloid nucleus, bed nucleus of the anterior commissure, striatum, accumbens nucleus, thalamic reticular nucleus, anteroventral and paraventricular thalamic nuclei, granular layer of the cerebellar cortex, anterior and ventral tegmental nuclei, granular layer of the cochlear nucleus, and the parvicellular part of the lateral reticular nucleus (Ohishi et al., 1998). In the cerebellar cortex, cell bodies and dendrites of about 90% of the total population of Golgi cells showed mGluR2 immunoreactivity; it was indicated that Golgi cells with mGluR2 were segregated from those with mGluR5 (Neki et al., 1996b). No particular species differences were found in the distribution pattern of mGluR2 immunoreactivity between rat and mouse (Ohishi et al., 1998).

2.3.5. mGluR3 immunoreactivity The distribution of mGluR3 in the brain and spinal cord was examined immunohistochemically in the mouse by using an antibody that was raised against a fusion protein containing 82

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amino acid residues 824-879 of rat mGluR3 (Table 2; Y. Tamaru et al., unpublished). Since this mGluR3 antibody somewhat cross-reacted with mGluR2, the results obtained from wild-type mouse were evaluated by comparing them with those obtained from the mGluR2 gene-lacking mouse (Yokoi et al., 1996). Immunoreactivity for mGluR3 was found extensively in the brain and spinal cord of the mouse. Intense immunoreactivity was observed in the olfactory tubercle, piriform cortex, neocortex, limbic cortex including the cingulate, retrosplenial, perirhinal, entorhinal and subicular cortical areas, CA3 stratum lacunosum, molecular layer of the dentate gyrus, lateral and basolateral amygdaloid nuclei, lateral septal nucleus, striatum, and accumbens nucleus; some part of the globus pallidus and the external part of the anterior olfactory nucleus also showed intense immunoreactivity. Thus, the distribution pattern of mGluR3 immunoreactivity was in good accordance with that of cell bodies that expressed mGluR3 mRNA (Ohishi et al., 1993b). Electron-microscopically, mGluR3 immunoreactivity was observed not only in postsynaptic elements but also in presynaptic elements and glial processes, as mGluR2/3 immunoreactivity. 2.4. DISTRIBUTION OF mRNA AND IMMUNOREACTIVITY FOR GROUP III METABOTROPIC GLUTAMATE RECEPTORS

2.4.1. mGluR4 mRNA The in situ hybridization histochemistry revealed a wide distribution of mGluR4 mRNA in the brain and spinal cord of the rat (Tanabe et al., 1993; Ohishi et al., 1995a). According to a systematic study (Ohishi et al., 1995a), expression of mGluR4 was most intense in the granule cells of the cerebellar cortex; prominent expression was also observed in the periglomerular cells and granule cells of the main olfactory bulb, olfactory tubercle, entorhinal cortex, hilus of the hippocampus, lateral septum, septofimbrial nucleus, the rostral part of the intercalated amygdaloid nucleus, thalamic nuclei, lateral mammillary nucleus, pontine nuclei, and spinal motoneurons. Expression of mGluR4 mRNA was also observed in CA2 (Fotuhi et al., 1994) and the striatum (Testa et al., 1994) of the rat; in the human brain, the expression of mGluR4 mRNA was reported in the striatum, thalamus, hypothalamus, and cerebellum; the strongest expression was seen in the granule cells in the cerebellar cortex (Makoff et al., 1996a). Distinct expression of mGluR4 mRNA was reported in motoneurons of the lumber spinal cord of the rat during development; this was significantly increased in the adult (Berthele et al., 1999). In the rat retina, expression of mGluR4 mRNA was observed in the cell bodies in the ganglion cell layer; these were presumed to be the retinal ganglion cells (Akazawa et al., 1994; Hartveit et al., 1995) and displaced amacrine cells (Hartveit et al., 1995). Expression was reported further in amacrine cells in the inner part of the inner nuclear layer (Hartveit et al., 1995).

2.4.2. mGluR4 immunoreactivity Intense mGluR4a immunoreactivity was found in the molecular layer of the cerebellar cortex of the rat; this was localized electron-microscopically in axon terminals of the parallel fibers arising from the granule cells in the cerebellar cortex (Kinoshita et al., 1996b; Mateos et al., 1999). Presynaptic localization of mGluR4a immunoreactivity was also reported in the other brain regions. In the trapezoid body of the rat, mGluR4a immunoreactivity was observed in 83

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axon terminals wrapping the principal globular neurons in the medial nucleus of the trapezoid body (Elezgarai et al., 1999). In the hippocampus of the rat, mGluR4a immunoreactivity was generally weak and diffuse, but it was moderate in the inner third of the molecular layer of the dentate gyms; presynaptic localization of the mGluR4a immunoreactivity was observed electron-microscopically in the inner third of the molecular layer of the dentate gyms, as well as in the stratum oriens of CA2 (Shigemoto et al., 1997). It was further reported in the rat and mouse that mGluR4a immunoreactivity was most intense in the molecular layer of the cerebellum, strong in the globus pallidus, and moderate in the substantia nigra pars reticulata and entopeduncular nucleus, and moderate to weak in the striatum neocortex, hippocampus, striatum, and thalamus; mGluR4a immunoreactivity in the globus pallidus was localized in axon terminals of striatopallidal fibers (Bradley et al., 1999). In the retina of the rat, mGluR4a immunoreactivity was observed throughout the entire inner plexiform layer, exclusively at the postsynaptic targets of the cone bipolar cells and the rod bipolar cells (Koulen et al., 1996). Immunoreactivity for mGluR4 (both mGluR4a and mGluR4b) was reported in non-pyramidal neurons in the cerebral cortex and in CA2 of the rat hippocampus (Phillips et al., 1997). 2.4.3. Distribution of mRNA and immunoreactivity for mGluR6

Blot and in situ hybridization analyses of the CNS of the rat indicated that expression of mGluR6 mRNA was restricted to the retina, and no obvious expression of mGluR6 mRNA was detected in any other regions of the brain (Nakajima et al., 1993). Expression of mGluR6 mRNA was seen in the outer part of the inner nuclear layer in the retina of the adult rat (Nakajima et al., 1993; Akazawa et al., 1994; Hartveit et al., 1995), and mGluR6 immunoreactivity was localized exclusively to the postsynaptic, dendritic part of rod bipolar cells in the adult rat retina (Nomura et al., 1994). Developmental changes in subcellular localization of mGluR6 immunoreactivity were also observed: labeling for mGluR6 was initially distributed diffusely in both the cell bodies and dendrites of the rod bipolar cells, but gradually became punctate in the outer plexiform layer during the second postnatal week and finally concentrated on the synaptic sites by postnatal day 28 (Nomura et al., 1994). 2.4.4. mGluR7 mRNA

Distribution of mGluR7 mRNA was observed in the rat CNS by in situ hybridization histochemistry (Okamoto et al., 1994; Saugstad et al., 1994; Kinzie et al., 1995; Ohishi et al., 1995a; Corti et al., 1998; Kosinski et al., 1999). According to a systematic study (Ohishi et al., 1995a), mGluR7 mRNA was expressed widely in the CNS of the rat (Table 1): prominent expression of mGluR7 mRNA was seen in the main and olfactory bulbs, olfactory tubercle, neocortex, limbic cortex including CA1-CA3 and dentate gyms, striatum, accumbens nucleus, claustrum, amygdaloid complex, preoptic region, hypothalamus, thalamus, Purkinje cells of the cerebellar cortex, many regions in the lower brainstem, and dorsal horn of the spinal cord. Most intense expression was seen in the tufted and mitral cells of the olfactory bulbs, medial septal nucleus neurons, and locus coeruleus. The ganglion neurons in the trigeminal and the dorsal root ganglia also were labeled intensely. In the lumber spinal cord of the neonatal rat, it was reported that mGluR7 mRNA was expressed relatively strongly in the dorsal horn with the highest density in laminae I and II, and weakly throughout the rest of the spinal cord; there was a tendency for a decrease in the 84

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expression in the dorsal horn with maturity while the motoneurons showed no alteration in expression (Berthele et al., 1999). It was reported that hybridization signals of mGluR7a were higher than those of mGluR7b in the neocortex, CA3 area of the hippocampus, anterior thalamus, medial geniculate nucleus, and locus coeruleus (Corti et al., 1998). In the retina of the rat, expression of mGluR7 mRNA was reported in the cells of the inner nuclear layer and ganglion cell layer; these cells were presumed to be the retinal ganglion cells (Akazawa et al., 1994; Hartveit et al., 1995), or to be the amacrine cells, and bipolar cells (Hartveit et al., 1995).

2.4.5. mGluR7 immunoreactivity Distributions of two alternative splicing variants of mGluR7, mGluR7a and mGluR7b, were examined systematically in the CNS of the rat and mouse by using variant-specific antibodies raised against C-terminal portions of rat mGluR7a and human mGluR7b (Kinoshita et al., 1998): the distribution pattern of the immunoreactivity was compatible with that of mGluR7 mRNA, although the distribution of mGluR7b immunoreactivity was more limited than that of mGluR7a immunoreactivity; many CNS regions showing mGluR7a immunoreactivity displayed no mGluR7b immunoreactivity, while most regions showing mGluR7b immunoreactivity also displayed mGluR7b (Table 2). It was also revealed that the distribution pattern in the rat was substantially the same as that in the mouse, although some species differences were observed in the medial habenular nucleus, cerebellar deep nuclei, and lateral vestibular nucleus. In the medial habenular nucleus, mGluR7a immunoreactivity was intense in the rat, but was hardly detectable in the mouse. In the cerebellar nuclei and the lateral vestibular nucleus, mGluR7b immunoreactivity was more marked in the mouse than in the rat. It was also reported that mGluR7a was widely distributed throughout the rat brain, with a high level of expression in sensory areas, such as the piriform cortex, superior colliculus, and dorsal cochlear nucleus (Bradley et al., 1998). Distribution of mGluR7a immunoreactivity was also studied in the rhinencephalic regions (Kinzie et al., 1997; Wada et al., 1998), hippocampus (Bradley et al., 1996; Shigemoto et al., 1996, 1997), and autonomic cell groups of the medulla oblongata (Hay et al., 1999). In the rat hippocampus (Shigemoto et al., 1997), immunoreactivity for both mGluR7a and mGluR7b was observed exclusively in axon terminals. Immunoreactivity for mGluR7a was seen in all dendritic layers throughout the hippocampus, while mGluR7b immunoreactivity was observed only in the terminal zone of the mossy fibers; virtually all mGluR7b immunoreactive structures were also mGluR7a immunoreactive. Localization of mGluR7a immunoreactivity in axon terminals was also observed in primary afferent fibers terminating in laminae I and II of the spinal dorsal horn of the rat (Ohishi et al., 1995b), in the islands of Calleja (Kinoshita et al., 1998), and in layer I of the piriform cortex of the rat (Kinzie et al., 1997; Wada et al., 1998). It was reported, however, that mGluR7 immunoreactivity was seen occasionally in somatodendritic domains of neurons in the hippocampus, locus coeruleus, cerebellum, and thalamic nuclei of the rat (Bradley et al., 1998). In the basal ganglia of the rat, Kosinski et al. (1999) confirmed the presence of mGluR7a immunoreactivity in axon terminals of corticostriatal, striatopallidal, and striatonigral fibers. They further reported the existence of mGluR7a immunoreactivity in dendrites and spines in the striatum and globus pallidus. In the retina of the rat, mGluR7a immunoreactivity was present exclusively in the inner plexiform layer (Brandst~itter et al., 1996). Electron-microscopy revealed immunoreactivity 85

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for mGluR7a presynaptically in OFF- and ON-cone bipolar cell ribbon synapses, and postsynaptically in amacrine cells or, in a very few cases, in ganglion cell dendrites. The presynaptic mGluR7a immunoreactivity was restricted to one half of the release site facing only one of the two postsynaptic processes, indicating differential regulation of glutamate release from the ribbon synapse to the postsynaptic neurons.

2.4.6. mGluR8 mRNA The expression pattern of mGluR8 mRNA was studied by in situ hybridization histochemistry in the mouse (Duvoisin et al., 1995) and rat (Saugstad et al., 1997; Corti et al., 1998); it showed highly restricted distribution as compared with that of mGluR4 and mGluR7. In the rat (Saugstad et al., 1997; Corti et al., 1998), prominent mGluR8 expression was observed in the mitral cell layer and granule cell layer of the main and accessory olfactory bulbs, piriform cortex, pontine nuclei, and lateral reticular nucleus of the medulla oblongata. In the main olfactory bulb, the expression was more intense in the mitral cell layer than in the granule cell layer, while it was more intense in the granule cell layer than in the mitral cell layer in the accessory olfactory bulb. Pyramidal cells of the piriform cortex also showed high levels of expression, whereas the olfactory tubercle was virtually lacking mGluR8 mRNA. Low levels of expression was also detected in the layers V and VI of the cerebral neocortex, hippocampus, septum, basolateral amygdaloid nuclear group, thalamic reticular nucleus, mammillary nuclei, and cerebellum. In general, hybridization signals of mGluR8a were higher than those of mGluR8b in the majority of the brain regions; in some areas, such as the spinal vestibular nucleus, ambiguus nucleus, and lateral nucleus of the medulla oblongata, only mGluR8a was detected (Corti et al., 1998). Some species differences appeared to exist in the expression pattern of mGluR8 mRNA between the rat and mouse: in the mouse (Duvoisin et al., 1995), strong expression was reported in the main and accessory olfactory bulbs including the granule, mitral and periglomerular layers, olfactory tubercle, and mammillary nuclei. Expression was also observed in scattered cells in the deeper layers of the cerebral cortex and in the hind brain, but not in the hippocampus and cerebellum; no particular description was given about the expression in the lateral reticular nucleus of the medulla oblongata, although a low level of expression was reported in the retina. The mGluR8 mRNA expression in the brain and retina was reported to be stronger and more widely spread in the developing mouse than in the adult (Duvoisin et al., 1995): it was observed with varying intensities in parts of the developing telencephalon, thalamus, hypothalamus, midbrain, pons, and medulla oblongata, as well as in the olfactory bulb and retina. In the developing retina, expression was seen in the ganglion cell and inner nuclear cell layers and possibly in the outer nuclear layer. Expression was also detected in the developing trigeminal and dorsal root ganglia.

2.4.7. mGluR8 immunoreactivity Expression mGluR8a immunoreactivity was examined in the rhinencephalic regions (Kinoshita et al., 1996a; Wada et al., 1998) and hippocampus (Shigemoto et al., 1997) of the rat: immunoreactivity for mGluR8a was observed in the external and internal plexiform layers of the main olfactory bulb, mitral cell layer of the accessory olfactory bulb, anterior olfactory nucleus, superficial layers of the olfactory tubercle and layer Ia of the piriform and entorhinal cortical regions. In the hippocampus, expression of mGluR8 immunoreactivity was rather 86

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weak and diffuse, but marked in terminal zone of the lateral perforant path, i.e., the outer layer of the CA3 stratum lacunosum-moleculare and the outer one third of the molecular layer of the dentate gyrus. Electron-microscopically, mGluR8 immunoreactivity was observed in axon terminals in layer Ia of the piriform cortex (Kinoshita et al., 1996a; Wada et al., 1998) and in the molecular layer of the dentate gyrus (Shigemoto et al., 1997).

3. DIFFERENTIAL SUBCELLULAR LOCALIZATION OF METABOTROPIC GLUTAMATE RECEPTORS IN RELATION TO TRANSMITTER RELEASE SITES As described in the previous sections, patterns of subcellular localization of mGluRs are different among three subgroups: group I and III mGluRs are mainly localized to somatodendritic and axonal domains of neurons, respectively, while group II mGluRs are extensively localized to both domains as well as to glial cell processes. In many cases, these observations were confirmed with pre-embedding immunoperoxidase electron-microscopy, the most sensitive method of immunolabeling. However, the difference of precise localization between mGluR subtypes within these domains is not readily resolved by this method because the peroxidase end-product may diffuse to membrane compartments without receptors (Luj~n et al., 1996). To avoid this problem, a high-resolution method with non-diffusible immunogold particles has been used to reveal differential localization of mGluRs targeted to specific membrane compartments in both postsynaptic and presynaptic elements. In some studies, quantitative analyses further revealed distinct patterns of immunoparticle distribution relative to glutamate release sites. 3.1. mGluRs IN POSTSYNAPTIC ELEMENTS Electron-microscopical immunogold detection of mGluRla immunoreactivity indicated that mGluRla was expressed preferentially at the periphery of the postsynaptic densities of asymmetrical synapses in the cerebellar cortex and hippocampus (Fig. 3; Baude et al., 1993; Nusser et al., 1994; Luj~n et al., 1996, 1997), as well as in the cortico(areal7)-thalamic synapses in the dorsal lateral geniculate nucleus, lateral posterior nucleus (Vidny~nszky et al., 1996), and ventral posterior thalamic nucleus (Liu et al., 1998). In symmetrical GABAergic synapses in the monkey pallidum, however, a large population of immunogold particles for mGluRla and mGluR5 were seen in the main body of the postsynaptic specializations (Hanson and Smith, 1999). At the heads of the spines of Purkinje cells in the rat cerebellar cortex, about half of the immunogold particles indicating mGluRla immunoreactivity were localized perisynaptically, i.e. within a 60-nm annulus surrounding the edge of the postsynaptic specialization, while the remaining particles were distributed extrasynaptically, i.e. at a more distant position, but not at the postsynaptic specialization (Luj~n et al., 1997). Immunogold labeling for mGluR5 was also observed preferentially at the periphery of the postsynaptic densities of asymmetrical synapses in the rat hippocampus (Luj~n et al., 1996, 1997). At the heads of the spines of CA1 pyramidal neurons, about one fourth of immunogold particles were localized perisynaptically, while the remaining particles were expressed extrasynaptically, but not at the postsynaptic specialization (Luj~n et al., 1997). Requirement of repetitive synaptic stimulation to detect activation of group I mGluRs (Batchelor et al., 1994; Congar et al., 1997; Fiorillo and Williams, 1998) may be ascribed to the extrasynaptic location of receptors (Baude et al., 1993; Nusser et al., 1994). The 87

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Fig. 3. Distinct subcellular localization of group I (mGluRla), group II (mGluR2/3) and group III (mGluR7b) mGluRs in relation to glutamate release sites. Pre-embedding immunogold method revealed typical labeling patterns for mGluRla, mGluR2/3, and mGluR7b in the molecular layer of the cerebellum, stratum lacunosum moleculare in the CA1 area of the hippocampus, and stratum lucidum in the CA3 area, respectively. Immunoparticles for mGluRla are mostly found in postsynaptic elements and concentrated around asymmetrical synapses (arrowheads) between parallel fiber terminals and Purkinje cell dendritic spines. Labelings for both mGluR2/3 and mGluR7b are found in presynaptic elements but with a distinct relation to asymmetrical synapses; immunoparticles for mGluR7b are concentrated in the presynaptic active zone, whereas those for mGluR2/3 are diffusely distributed on membranes remote from the active zone in preterminal axons and axon terminals. Scale bar is 0.5 I~m.

perisynaptic position of group I mGluRs is also consistent with the recent discovery of synapse associating proteins (Tu et al., 1999) linking group I mGluRs and NMDA receptors located in postsynaptic density, and membrane-delimited modulation of NMDA currents by mGluR1/5 in cultured cortical neurons (Yu et al., 1997). In contrast with the perisynaptic localization of group I mGluRs, mGluR2 immunogold labeling in the cerebellar Golgi cell dendrites showed no close association with glutamatergic synapses between Golgi cell dendrites and parallel fiber terminals (Luj~.n et al., 1997). Immunoparticles for mGluR2, however, occur in clusters in extrasynaptic sites (Luj~n et al., 1997), which may reflect association with other kinds of associating molecules in a specific membrane compartment. Another member of group II mGluRs, mGluR3, is also observed in postsynaptic elements in various brain regions (Y. Tamaru et al., unpublished). However, analysis of mGluR3 immunoparticles in the dentate molecular layer revealed a distribution pattern quite different from that of mGluR2 in Golgi cell dendrites: in the heads of the dendritic spines, about 20% of immunogold particles were seen in postsynaptic membrane specialization and 40% in perisynaptic plasma membranes within 60 nm from the edge of asymmetrical synapses (Y. Tamaru et al., unpublished). Thus, mGluR3 was indicated to be associated even more closely to glutamatergic synapses than group I mGluRs in the dentate gyrus. It is not yet clear if the difference between the location of mGluR2 and mGluR3 relative to glutamate release sites depends on subtypes or cell types. 3.2. mGluRs IN PRESYNAPTIC ELEMENTS In axon and axon terminals, group II and group III mGluRs are distributed differentially relative to neurotransmitter release sites (Fig. 3; Shigemoto et al., 1997; Wada et al., 1998). Group III receptors are mainly localized to the presynaptic active zone whereas group II receptors are often observed in extrasynaptic sites remote from the active zone in preterminal portions of axons and axon terminals. About 79% of immunoparticles for mGluR2 in cerebellar Golgi cell axons and about 72% of those for mGluR3 in corticostriatal axons were 88

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found in extrasynaptic membranes and only 2-4% of those were detected in the presynaptic active zone (Lujfin et al., 1997; Y. Tamaru et al., unpublished). In contrast, the majority of the immunogold labeling for mGluR4, mGluR7a, mGluR7b, and mGluR8a was detected in the presynaptic active zone in glutamatergic terminals in various brain regions (Shigemoto et al., 1996, 1997; Wada et al., 1998; Mateos et al., 1999). This distinct segregation of group II and group III mGluRs may indicate different sources of glutamate activating these receptors and different effector molecules coupled to these receptors. It is conceivable that group III mGluRs function as autoreceptors activated by glutamate released from the active zone where they are located, whereas group II mGluRs might be activated by spillover glutamate from distant synapses on the same or other presynaptic elements (Vogt and Nicoll, 1999). In the cerebellar cortex of the rat, it is suggested that mGluR2/3 on the axonal domain of Golgi cells might mediate heterosynaptic inhibition from the adjacent mossy fiber terminals, i.e. transmitter glutamate released from the mossy fibers terminals might activate mGluR2/3 on axon terminals of Golgi cells to control the GABA release from the axon terminals of Golgi cells (Ohishi et al., 1994). Immunogold particles for mGluR3 were also found in GABAergic projection fibers in the thalamus suggesting heterosynaptic sources of glutamate activating these receptors (Y. Tamaru et al., unpublished). Furthermore, group III mGluRs are also present in some GABAergic terminals (Kinoshita et al., 1998; Bradley et al., 1999) and heterosynaptic interaction should be taken into account for these receptors as well. Presynaptic inhibition of transmission mediated by group II and group III mGluRs has been reported very widely in the brain with diverse mechanisms such as suppression of presynaptic voltage-dependent calcium channels, activation of presynaptic K channels, and direct inhibition of exocytosis (for review, see Anwyl, 1999). However, similar effector mechanisms are reported with agonists selective for group II and group III mGluRs and the functional difference corresponding to distinct presynaptic localizations of these receptors remains elusive. In addition, much convincing evidence for presynaptic inhibitory effects mediated by group I mGluRs is reported but there has been very little corresponding morphological evidence for presynaptic mGluR1 and mGluR5 except that in peculiar dendrodendritic synapses in the olfactory bulb (Van den Pol, 1995). This discrepancy may be due to the limit of sensitivity for detecting immunoreactivity for group I mGluRs. However, it should also be noted that activation of postsynaptic mGluRs might be involved in the expression of presynaptic inhibition through retrograde signalling mechanisms (Harvey et al., 1996). 3.3. TARGET-CELL-SPECIFIC SEGREGATION OF GROUP III mGluRs One of the most peculiar findings on presynaptic mGluR localization is target-cell-specific concentration of group III mGluRs in the presynaptic active zone (Fig. 4; Shigemoto et al., 1996, 1997). In the rat hippocampus, pyramidal cell axon terminals presynaptic to a particular subpopulation of GABAergic interneurons (somatostatin/mGluRla-positive cells) have a much higher level of presynaptic mGluR7a than axon terminals making synapses with pyramidal cells and other types of interneurons. Synapses emanating from the same axon, even within the same terminals, exhibited different densities of mGluR7a, depending on the nature of the postsynaptic target (Shigemoto et al., 1996). The segregation of mGluR7a between two release sites of a single terminal implies that coupling of the receptor to its effector molecules is spatially restricted and probably membrane-delimited to ensure the specificity of local regulation. The similar target-specific segregation was also found for other subtypes of group III mGluRs (Shigemoto et al., 1997) and in other brain regions and the retina 89

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.

.

.

i.,% ~

Fig. 4. Target-cell-specific segregation of group III mGluRs in the presynaptic active zone. Pre-embedding immunogold labeling for mGluR7a in the CA3 area of the hippocampus and that for mGluR8a in the layer Ia of the piriform cortex in the rat are shown. Single terminals (tl and t2) make two asymmetrical synapses but only presynaptic active zones making contacts on dendritic shafts (d) are heavily labeled with immunoparticles whereas those on spines (s) have no or very weak labeling. The target dendrites (d) make many other asymmetrical synaptic contacts with terminals heavily labeled for respective mGluRs. Scale bar is 0.5 I~m.

(Fig. 4; Brandst~itter et al., 1996). These findings imply that postsynaptic cells could influence the regulation of transmitter release by controlling the density of presynaptic receptors in the active zone through some retrograde signaling mechanisms. Indeed, target-cell-specific regulation of presynaptic short-term and long-term plasticity have been reported in neocortical pyramidal cell axons (Reyes et al., 1998) and hippocampal mossy fibers (Maccaferri et al., 1998), respectively. In the former, facilitation of excitatory postsynaptic potentials (EPSPs) was observed in somatostatin-positive interneurons by repetitive action potentials induced in pyramidal cells while the same stimulation showed depression of EPSPs in parvalbumin-positive interneurons. Although somatostatin-positive interneurons seem to be decorated with terminals intensely labeled for mGluR7a, the functional link between mGluR7a and facilitation of EPSP remains unclear. Anyway, to understand the physiological roles of mGluR-mediated regulation of synaptic transmission, it is thus very important to determine the identity of postsynaptic targets in addition to that of presynaptic terminals with group III mGluRs.

4. ABBREVIATIONS

Acb AOB Cb CNS Cx DCG-IV DG DHPG EPSPs GP Hi IC 90

nucleus accumbens accessory olfactory bulb cerebellum central nervous system neocortex (2S, l'R,2'R,3'R)-2-(2,3-dicarboxycyclopropyl) glycine dentate gyrus 3,5-dihydroxyphenylglycine excitatory postsynaptic potentials globus pallidus hippocampus inferior colliculus

Metabotropic glutamate receptors L-AP4 LS LY354740 mGluRs MOB OT Pit Rt SC SN SpV St Th VP

Ch. III

L-2-amino-4-phosphonobutyrate lateral septum (+)- 1S,2S,5R,6S-2-aminobicyclo[3.1.0]hexane-2,6-dicarboxylic acid metabotropic glutamate receptors main olfactory bulb olfactory tubercle piriform cortex reticular thalamic nucleus superior colliculus substantia nigra spinal trigeminal nucleus caudal part neostriatum thalamus ventral pallidum

5. ACKNOWLEDGEMENTS The authors wish to thank A. Kinoshita for critically reading the manuscript, Y. Tamaru for unpublished data, and H. Kuzume and S. Doi for technical assistance. This work was supported by grants from the Ministry of Education, Science and Culture of Japan (R.S., N.M.) and CREST of Japan Science and Technology Corporation (R.S.).

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CHAPTER IV

AMPA, kainate and NMDA ionotropic glutamate receptor expression an in situ hybridization atlas W. WISDEN, EH. SEEBURG AND H. MONYER

1. INTRODUCTION The ionotropic glutamate receptors are classified into NMDA, AMPA and kainate subtypes (Dingledine et al., 1999; Hollmann, 1999). The receptors are heteromeric assemblies of different subunits. Knowing how this large family of subunit genes is expressed in the brain tells us which neural circuits and systems they contribute to. In this chapter, we review the distribution of the rodent NMDA, AMPA and kainate receptor subunit mRNAs as mapped by in situ hybridization (ISH). In the final analysis it is essential to know where the proteins are located on the cell (Petralia, 1997; Somogyi et al., 1998), but mRNA distributions give us a reliable picture of how a gene family is expressed; there are no problems of antibody specificity and cross-reactivity. The glutamate receptor subunit genes are expressed in all areas of the central and peripheral nervous system (Wisden and Seeburg, 1993b; Bahn and Wisden, 1997; Watanabe, 1997), and also in non-neuronal lineages. In this chapter, we describe in detail the distribution of receptor subunit mRNAs in the retina, the neocortex, the hippocampus, the striatum (caudate putamen), the cerebellum, and the spinal cord (Table 1). These brain areas exemplify complex circuits using multiple receptor subtypes.

2. AMPA AND KAINATE RECEPTORS

The non-NMDA ionotropic receptors consist of two subgroups: AMPA and kainate receptors (Dingledine et al., 1999; Hollmann, 1999). In the CNS, AMPA receptors are responsible for 'general purpose' excitatory transmission at most synapses (reviewed in: Geiger et al., 1999; Monyer et al., 1999); their properties allow high temporal precision, short latency of action potential initiation, and EPSP coincidence detection. Kainate receptor function, on the other hand, is subtle and not understood (Lerma, 1999). Kainate receptor responses are small. At glutamatergic synapses of hippocampal CA1 neurons, the peak synaptic response of the kainate receptor-activated current is less than a tenth of the AMPA receptor response, but lasts longer (reviewed in: Geiger et al., 1999; Lerma, 1999). The slow excitation initiated by kainate receptors may integrate synaptic activity and/or adjust the membrane potential close to the threshold for action potential initiation (Geiger et al., 1999). Functional kainate receptors have been demonstrated on hippocampal GABAergic interneurons (Cossart Handbook of Chemical Neuroanatomy, Vol. 18: Glutamate O.P. Ottersen and J. Storm-Mathisen, editors (~ 2000 Elsevier Science B.V. All rights reserved.

99

TABLE 1. Expression of NMDA, AMPA and kainate receptor subunit mRNAs in selected cell types of the rat central nervous system Hippocampus

Cerebellum

Dentate granule cell

CA1 pyramidal cell

CA3 pyramidal cell

Purkinje cell

GluR-Ai,o

GluR-Ao GluR-Ai (lower) GluR-Bo GIuR-Bi (lower) GluR-Ci,o GluR-Do

GluR-Ai GluR-Ao (lower) GluR-Bi GluR-Bo (lower) GluR-Ci

GluR-Ao

GluR-Bi,o GluR-Ci,o GluR-Do KA1 KA2

GluR-Bi,o

Granule cell cell

KA2

GluR6 GluR7

GluR6

GluR6

NR 1 NR2A NR2B

NR 1 NR2A NR2B

NR 1 NR2A NR2B

Caudate putamen

Motor neuron

Medium spiny neuron

Cholinergic interneuron

Retinal OFF-bipolar cell

GluR-Ai,o

GluR-A

GluR-A

GluR-Ai GluR-Bi

Retina

GluR-B

GluR-Bi,o

GluR-Bi,o

GluR-Ci,o GluR-Di

GluR-Ci (lower) GluR-Do (lower)

GluR-D

KA2

KA2

KA2

KA2

GluR6

GluR6 GluR7

GluR6

GluR5 GluR6 GluR7

NR 1 NR2A NR2B

NR 1

NR1

GluR-Ci GluR-Do

KA1 KA2

Bergmann glia

Spinal cord

GluR-Di

KA1

KA1

GluR5

NR 1

NR 1 NR2A

NR 1 NR2B

NR 1 NR2A NR2B (low)

NR2B NR2C

NR2C NR2D (low) NR3A (low)

NR2D (low) NR3A (low) 1 (low)

~ 1 (low)

~ 1 (low)

NR2D

~ 1 (low) ~2

Flip and flop splice forms of AMPA receptor subunits are indicated by i and o suffixes. Modified from Wisden and Seeburg (1993b). Glutamate receptor subunit mRNAs in selected adult rat CNS cells.

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et al., 1998; Frerking et al., 1998), and on hippocampal pyramidal cells (Bureau et al., 1999). 2.1. AMPA RECEPTOR SUBUNITS m SUMMARY OF mRNA DISTRIBUTION AMPA receptors are heteromers of the GluR-A to -D (GluR-1 to -4) subunits. If the GluR-B subunit is in the complex, the receptor is less CaZ+-permeable, and has a two- to three-fold lower single-channel conductance (Swanson et al., 1997; Monyer et al., 1999). Receptors with the GluR-D subunit have faster kinetics. Each AMPA receptor subunit exists as either a flip or a flop variant, determined by mutually exclusive splicing of an exon encoding a domain of 38 amino acids (Sommer et al., 1990). This domain influences receptor desensitization (Sommer et al., 1990; Mosbacher et al., 1994). The overall distribution of AMPA receptor subunit mRNAs is shown in Fig. 1 (Boulter et al., 1990; Kein~inen et al., 1990; Gold et al., 1997). GluR-A mRNA is most abundant in the hippocampus, amygdala and cerebellar Bergmann glia. GluR-B is nearly universally expressed, but its expression is particularly high in cerebellar granule cells, neocortex and the hippocampus. GluR-B is absent or expressed at lower levels in most GABAergic interneuron types (Monyer et al., 1999). GluR-C expression is highest in neocortex and hippocampus. GluR-D expression is highest in the cerebellum (granule cells and Bergmann glial cells) with comparatively light expression in the forebrain. Expression of all four AMPA receptor subunits is prominent in the olfactory bulb and medial habenula (Boulter et al., 1990; Kein~inen et al., 1990). Curiously, relative to other areas, there is little AMPA receptor subunit mRNA or protein in the thalamus, with the exception of GluR-D in the reticular thalamic nucleus (Kein~inen et al., 1990; Gold et al., 1997). Throughout the brain, the flip and flop splice forms have different expressions (Fig. 2); this is well illustrated by looking at the hippocampus (see Section 7.2.1.1) (Sommer et al., 1990). As outlined above, GABAergic interneurons express GluR-B subunit mRNA and protein at lower levels than principal neurons (reviewed in: Petralia et al., 1997; Geiger et al., 1999); the flop splice forms predominate in interneurons, and some interneuronal types strongly express GluR-D (Geiger et al., 1999). The low GluR-B flip content and high GluR-D expression in GABAergic interneurons may be responsible for the rapid AMPA receptor deactivation of interneurons (Monyer et al., 1999). 2.2. KAINATE AND 8 RECEPTOR SUBUNITS - - SUMMARY OF mRNA DISTRIBUTION Kainate receptors are heteromeric and homomeric assemblies of GluR5, GluR6, GluR7, KA1 and KA2 (Herb et al., 1992; Cui and Mayer, 1999; Dingledine et al., 1999; Paternain et al., 2000). The 8 subunits might assemble with either AMPA or kainate receptor subunits. In adult brain, 81 expression is low; its highest mRNA levels are in the hippocampus (Fig. 3) (Yamazaki et al., 1992; Lomeli et al., 1993); 82, in contrast, is highly expressed in cerebellar Purkinje cells, with low expression elsewhere (Fig. 3) (Araki et al., 1993; Lomeli et al., 1993). The most significant site in the brain where the 82 subunit makes a contribution is the parallel fibre synapse on the cerebellar Purkinje cell (Zhao et al., 1997); loss and gain of 82 gene function causes Purkinje cell malfunction (no LTD at the granule cell-Purkinje cell synapses) and death, respectively (Kashiwabuchi et al., 1995; Zuo et al., 1997). As adult Purkinje cells contain no functional NMDA receptors (Cull-Candy et al., 1998), the 82 subunit probably contributes to AMPA or kainate receptors or to an unknown GluR type. 101

Ch.

IV

W. W i s d e n

B

e t al.

~

CAt.

H ~j

.,~

'}

La

Fig. 1. AMPA receptor subunit mRNA dist

.,o~i

R

........

........... /

=tion f in horizontal and coronal sections of adult rat brain. (A and E), GluR-A expression; (B and F), GluR-B; (C and G), GluR-C; (D and H), GluR-D. Cb, cerebellum; CIC, central nucleus of the inferior colliculus; CPu, caudate putamen; Cx, neocortex; DG, dentate gyms; E, entorhinal cortex; La, lateral amygdaloid nucleus; Me, medial amygdaloid nucleus; OB, olfactory bulb; Rt, reticular thalamic nucleus; S, septal nuclei; SC, superior colliculus (deep layers); VM, ventral medial thalamic nucleus; Scale bar H, 3.7 mm. (l-L), AMPA receptor subunit mRNA distribution in adult rat cerebellum. (/) GluR-A distribution; arrowheads mark the line of silver grains along the Purkinje-Bergmann glia cell layer; this is due to GluR-A expression in Bergmann glia; (J) GluR-B, arrowheads mark labelled Purkinje cells; (K) GluR-C, unlabelled arrowheads mark silver grain clusters in the molecular layer over stellate/basket cells; (L), GluR-D, arrowheads as in I. gr, cerebellar granule cell layer; mol, molecular layer; P, Purkinje cells; wm, white matter. Scale bar L, 500 Ixm (Kein~inen et al., 1990).

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ob

Fig. 2. Distribution of AMPA receptor flip and flop splice variant mRNAs in the adult rat brain (X-ray film autoradiographs, sagittal sections), cb, cerebellum" cpu, caudate putamen; ctx, neocortex" dg, dentate granule cells; ob, olfactory bulb (Sommer et al., 1990; Wisden, Seeburg and Monyer, unpublished). Scale bar, 10 ram.

An example of kainate receptor subunit mRNA distribution in the rat brain is shown in Fig. 4 (Wisden and Seeburg, 1993a). The 'fingerprint' of KA1 expression is hippocampal CA3 pyramidal cells and dentate granule cells (Werner et al., 1991), whereas KA2 is expressed at moderate levels throughout the brain (Herb et al., 1992). KA1 mRNA is also found in glial cells, in the corpus callosum and cerebellar white matter tracts (Wisden and Seeburg, 1993a). GluR5 mRNA is most abundant in cerebellar Purkinje cells, the cingulate and piriform cortex, several septal, thalamic and hypothalamic nuclei, and the amygdala (Bettler et al., 1990; Wisden and Seeburg, 1993a; Bahn et al., 1994); it is possible that GluR5 is mainly expressed in GABAergic cells. In both mouse and rat, GluR6 mRNA levels are highest in cerebellar granule cells, and there is moderate GIuR6 expression in the hippocampus and caudate putamen (Egebjerg et al., 1991; Wisden and Seeburg, 1993a; Bahn et al., 1994). The GluR7 gene is expressed mainly in the deep layers of neocortex, reticular thalamic nucleus and the cerebellar stellate/basket cells (Bettler et al., 1992; Lomeli et al., 1992; Wisden and Seeburg, 1993a; Bahn et al., 1994). 103

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om

Fig. 3. Distribution of 31 (A) and 32 (B) subunit mRNA in the adult rat brain (X-ray film autoradiographs, horizontal sections); (C), localization of 32 mRNA in cerebellar Purkinje cells (emulsion autoradiograph). Cb, cerebellum; Ctx, neocortex; Gr, granule cells; H, hippocampus; Mol, molecular layer; P, Purkinje cells; arrowheads mark labelled Purkinje cells. Scale bar in B, 3.5 mm; scale bar in C, 28 I~m (Lomeli et al., 1993).

3. NMDA RECEPTORS NMDA receptors have a voltage-dependent Mg 2+ block. This means that they open to glutamate only when the membrane in which they sit is already depolarized. They stay open much longer than AMPA receptors (hundreds of milliseconds rather than milliseconds), and are highly CaZ+-permeable (reviewed in: Bliss and Collingridge, 1993; Spruston et al., 1995; Monyer et al., 1999). These integrative properties make NMDA receptors essential for many 104

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Fig. 4. (A-J) Distribution of kainate receptor subunit mRNAs in the adult rat brain (X-ray film autoradiographs, coronal sections). Arrowheads in E and F mark neocortical layer III cells expressing the GluR7 gene. AV, anteroventral thalamic nucleus; BST, bed nucleus stria terminalis; CC, corpus callosum white matter tract; Cg, cingulate cortex; Cpu, caudate putamen; DG, denate granule cells; DM, dorsomedial hypothalamic nucleus; GP, globus pallidus; MPA, medial preoptic area; Pit, piriform cortex; Rt, reticular thalamic nucleus; SCh, suprachiasmatic nucleus. Scale bar, 3.2 mm (Wisden and Seeburg, 1993a).

types of synaptic plasticity including those involved in memory formation, the regulation of movement, and in influencing the sensory field size of receptive neurons in e.g. the spinal cord and visual cortex. 105

Ch. I V

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'

Fig. 5. The expression of the NMDA receptor subunit mRNAs (NR1, NR2A-NR2D) in the adult rat brain (X-ray

film autoradiographs, horizontal sections); (Monyeret al., 1992, 1994). 3.1. NMDA RECEPTOR SUBUNITS

SUMMARY OF mRNA DISTRIBUTION

The NMDA receptor subunit genes are NR1 (g 1), NR2A (el), NR2B (e2), NR2C (e3), NR2D (e4), NR3A (X-1 or NMDAR-L), and NR3B (Hollmann, 1999). The NR3B subunit is known only from partial genomic sequence (Hollmann, 1999). Most NMDA receptors are heteromeric NR1 and NR2 subunit assemblies (e.g. NR1/NR2A or NR1/NR2C). NR1 is a universal subunit, forming part of all NMDA receptors; the NR2 series affects the channel open time, channel conductance and Mg 2+ sensitivity (Monyer et al., 1994, 1999); the NR3A subunit reduces the single-channel conductance of NR1/NR2 complexes, and may be used to 'restrain' NMDA receptor function, particularly during development (Das et al., 1998). All the genes have different expression patterns (Kutsuwada et al., 1992; Monyer et al., 1992, 1994; Watanabe et al., 1993; Ciabarra et al., 1995; Laurie et al., 1997; Watanabe, 1997). The overall distributions of the NMDA receptor subunit mRNAs (NR1, NR2A-D) are shown in Fig. 5. The NR1 gene is expressed in most neuronal types (Moriyoshi et al., 1991); in some cells it is 106

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Fig. 6. Alternative splicing of the NMDA receptor NR1 subunit mRNA produces eight versions of the NR1 protein. The alternatively spliced cassettes are N1, C1 and C2. The nomenclature is given in the table. (Adapted from Zukin and Bennett, 1995.)

the sole NMDA receptor subunit gene expressed, e.g. in adult cerebellar Purkinje cells, retinal horizontal cells and spinal cord visceral motor neurons (Brandst~itter et al., 1994; Cull-Candy et al., 1998; Shibata et al., 1999).

3.1.1. NR1 RNA splice variants There are eight splice variants of the NR1 mRNA: one N-terminal exon insertion ( ' N I ' cassette: - e x o n 5 = NRI-a; +exon 5 = NRI-b), and seven C-terminal exon deletions [exon 21 encodes the C1 cassette, C1 deletion = NR1-2; exon 22 encodes the C2 cassette, C2 deletion -- NR1-3; combined C1 and C2 deletion = NR1-4 (reviewed by: Zukin and Bennett, 1995; Dingledine et al., 1999; Winkler et al., 1999) see Fig. 6]. The N1 cassette (exon 5) insertion influences gating kinetics. The C-terminal versions may regulate subcellular targeting and receptor clustering (reviewed in Dingledine et al., 1999). The splice variants are 107

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Fig. 7. The distribution of the NMDA receptor NR1 subunit mRNA splice variants in the adult rat brain (X-ray film autoradiographs, horizontal sections). Pan, NR1 total mRNA; AV, anteroventral thalamic nuclei; Cb, cerebellum; Cp, caudate putamen; Cx, neocortex; Dg, denate granule cells; ER, entorhinal cortex; Hi, hippocampus; S, septum; smc, sensori-motor cortex; T, thalamus. Scale bar, 1.8. mm (Laurie and Seeburg, 1994). See Fig. 6 for explanation of the nomenclature.

differentially distributed (Fig. 7) (Luque et al., 1994; Laurie and Seeburg, 1994; Laurie et al., 1995; Landwehrmeyer et al., 1995; T611e et al., 1995a; Paupard et al., 1997; Winkler et al., 1999): NRI-a expression is universal; NRI-b is widespread, but has little expression in the caudate putamen; and in the hippocampus, the expression is mainly in CA3 pyramidal cells 108

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Fig. 8. NMDA receptor subunit gene expression in the developing P7 postnatal brain (X-ray film autoradiographs) (Monyer et al., 1994). (Fig. 7). Sensori-motor cortex has enriched expression of the exon 5 insertion (Laurie and Seeburg, 1994) (Fig. 7). 3.1.2. The NR2 subunits

NR2A mRNA is widely expressed in the adult brain; NR2B mRNA is mainly forebrainspecific hippocampus and neocortex (Monyer et al., 1992, 1994; Fig. 5). NR2A gene expression increases strongly during postnatal development in many brains regions, e.g. neocortex, hippocampus and cerebellar granule cells (Monyer et al., 1994; Nase et al., 1999; Fig. 8). In contrast, during the same postnatal periods, NR2B expression stays either constant, increases but less so than that of NR2A, or decreases (e.g. in cerebellar granule cells) (Fig. 8). The NR2A/NR2B subunit ratio determines how long the receptors stay open receptors which contain more NR2B subunit stay open longer and may govern the LTD/LTP ratio at any given synapse, with receptors with NR2B promoting LTP induction (Tang et al., 1999). 109

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Fig. 9. The distribution of NR3A mRNA in the adult rat brain (X-ray film autoradiographs, coronal sections), am, amygdala; cm, centromedial thalamic nucleus; hy, hypothalamus; Co, superior colliculus; mg, medial geniculate nucleus; pn, pontine nucleus; pv, paraventricular thalamic nucleus; rt, reticular thalamus. Scale bar, 1 mm (Ciabarra et al., 1995).

The NR2C gene has its highest expression in cerebellar granule cells, with lower levels in the thalamus (Fig. 5). In many brain regions, including the neocortex, caudate putamen and spinal cord, the NR2C gene is weakly expressed in glial-like cells (Standaert et al., 1996, 1999; Shibata et al., 1999). NR2D mRNA is mainly expressed in the globus pallidus, thalamus, brainstem, and many GABAergic interneuron subtypes (Fig. 5) (Monyer et al., 1994; Standaert et al., 1996, 1999). The NR2D gene's highest expression level is during the early postnatal period (Fig. 8) (Watanabe et al., 1993; Monyer et al., 1994). 3.1.3. The NR3A subunit

Similar to NR2D, the NR3A gene's highest expression is during embryogenesis and the early postnatal period (Ciabarra et al., 1995; Sucher et al., 1995). Receptors with NR3A may contribute to shaping the dendritic tree (Das et al., 1998); however, in the adult, NR3A mRNA is found only in a few thalamic nuclei (paraventricular, centromedial, intermediodorsal, medial geniculate), the amygdala (Ciabarra et al., 1995; Sucher et al., 1995), the CA1 pyramidal cells of the hippocampus, and numerous scattered cells in the neocortex (Ciabarra et al., 1995; Fig. 9). This unusual expression pattern does not match any of the other NMDA receptor subunit genes, so presumably in the adult brain the NR3 subunit contributes to a very specific subset of NMDA receptors. There are two NR3A splice variants: NR3-short and NR3-1ong 110

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(Sun et al., 1998). NR3-1ong has a 20 amino acid insertion in the C-terminus; both forms are expressed in the adult; by RT-PCR, NR3-1ong expression is enriched in the cerebellum (Sun et al., 1998), but ISH with a probe recognizing both splice variants does not detect any cerebellar signal (Ciabarra et al., 1995).

4. RNA EDITING

The distribution of receptor subunit complexity made by RNA editing (Seeburg et al., 1998) cannot be mapped using ISH: just one or two nucleotides distinguish the edited and non-edited forms; hybridization probes do not cleanly distinguish between them. To map the distribution of edited mRNAs, RT-PCR has been done on brain region-specific cDNA or cDNA isolated from individual cells (Jonas et al., 1994; Monyer et al., 1999). These methods show that subunit microheterogeneity varies with brain region and cell type, e.g. for the AMPA receptor subunits, the flip or flop splice cassettes combine with the alternative versions of the R/G site-edited position (Lomeli et al., 1994). Editing of kainate receptor subunit mRNAs also varies with brain region (Seeburg et al., 1998). NMDA receptor subunit mRNAs are not edited.

5. RETINA

Glutamate's role as a neurotransmitter in the vertebrate retina is reviewed by Barnstable (1993), Brandst~itter et al. (1998) and Lo et al. (1998). As the cell bodies of different retinal cell types are in different laminae (Fig. 10), we can assign which general cell types express which glutamate receptor subunits. However, there are different subsets of the same cell class, e.g., there are at least 10 different types of on- and off-bipolars, and multiple subtypes of the other cell classes (Stevens, 1998). Without cell-type markers and double-labelling studies, ISH can not differentiate these. The cones and rods release glutamate onto the bipolar cells: only off-bipolars use ionotropic receptors at this synapse; on-bipolars use the metabotropic receptor mGluR6 instead. The distribution of NMDA and non-NMDA receptor mRNAs in the retina is summarized in Fig. 10. 5.1. NMDA RECEPTOR SUBUNIT mRNAs IN THE RETINA The NR1 gene is expressed in every neuronal type in the rat retina; the NR2A, 2B and 2C genes are expressed in different cell subsets (Fig. 10) (Brandst~itter et al., 1994); the NR2D gene is not expressed at all (Brandst~itter et al., 1994). Horizontal cells only have NR1 mRNA. At least six of the NR1 splice variants (see Fig. 6) are in the mouse retina as measured by RT-PCR: NRI-a and NRI-b (exon-5-containing), NR1-4 (no C1 or C2), NR1-3 (C1 only), NRI-1 (C1 plus C2), and NR1-2 (C2 only) (Lo et al., 1998). In addition to NR1, bipolar cells contain NR2C mRNA; NMDA receptors on these cells should have a lower degree of MgZ+-dependent voltage block (Brandst~itter et al., 1994). Labelling of the amacrine cell layer is 'patchy' with NR2A-, 2B- and 2C-specific probes; so probably subsets of amacrine cells express NR1/NR2A, NR1/2B or NR1/2C receptors, and some amacrine cells may have only NR1 mRNA (Brandst~itter et al., 1994). All ganglion cells are likely to express NMDA receptors, as all cells contain NR1, NR2A, NR2B and NR2C mRNAs (Brandst~itter et al., 1994; Hartveit et al., 1994). These could assemble as channels of high (NR1/NR2A/NR2B) and low (NR1/NR2C) conductance, and high and low Mg 2+ sensitivity, respectively. These receptors 111

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Fig. 10. The cell types in the rat retina and their expression of the AMPA, NMDA and kainate receptor subunit mRNAs (circuit diagram adapted from: Barnstable, 1993; Bahn and Wisden, 1997). The assignment of subunit groupings to different cell classes does not imply that, for example, all amacrine cells co-express all the listed subunits; subsets of amacrine cells, horizontal or ganglion cells express different subunit combinations (see text).

are likely to be post-synaptic to the glutamatergic bipolar cell terminals (Hartveit et al., 1994) (Fig. 10). This is a similar situation to adult cerebellar granule cells (Cull-Candy et al., 1998). 5.2. AMPA RECEPTOR SUBUNIT mRNAs IN THE RETINA The GluR-B gene expresses in every neuronal type of the rat retina; the other AMPA receptor subunit genes express in cell subsets (Hughes et al., 1992; Mtiller et al., 1992a; Hamasssaki-Britto et al., 1993; Fig. 10). Rat horizontal cells express the GluR-A, -B and -D subunit genes. In the cat retina, horizontal cells also strongly express the GluR-C gene, and so this is a species difference (Hamasssaki-Britto et al., 1993). The GluR-A and GluR-B genes express in bipolar cells (Hughes et al., 1992; Mtiller et al., 1992a), including (for the GluR-B) 112

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those bipolar cells associated with rods (from ISH on dissociated cells; Hughes et al., 1992). All four AMPA receptor subunit genes (GluR-A through to GluR-D) express in amacrine cells. The GluR-A, GluR-C and GluR-D genes express in ganglion cell subsets, suggesting that A/B, B/C and B/D heteromeric assemblies might be found. By hybridizing serial sections, GluR-A, -B and -C transcripts were found in single cat ganglion cells. Some ganglion cells had only two subunit types, but all cells had the GluR-B mRNA (Hamasssaki-Britto et al., 1993). Some young (P5) rat retinal ganglion cells have CaZ+-permeable AMPA receptors, although the receptors on most ganglion cells behave as if GluR-B is present (Taschenberger and Grantyn, 1998). Because of the near universal expression of GluR-B, most AMPA receptors in the retina will be Ca2+-impermeable and of low conductance. However, there could be multiple receptor subunit combinations on the same cells resulting in mixtures of CaZ+-permeable and Ca2+-impermeable receptors, as found for instance in hippocampal GABAergic interneurons (Toth and McBain, 1998) and pyramidal cells (Yin et al., 1999), cerebellar interneurons (Liu and Cull-Candy, 2000), as well as spinal cord motor neurons (Greig et al., 2000; Vandenberghe et al., 2000). 5.3. KAINATE RECEPTOR SUBUNIT mRNAs IN THE RETINA GluR5 mRNA is mainly in the somata of the outer two thirds of the inner nuclear layer (indicating expression in bipolar cells and horizontal cells) (Fig. 10). There are occasional GluR5-positive patches in the ganglion cell layer (Hughes et al., 1992; Mtiller et al., 1992a; Hamasssaki-Britto et al., 1993), implying expression in ganglion cell subsets or displaced amacrine cells (Mtiller et al., 1992a). The GluR6 gene expresses in subsets of amacrine cells and ganglion cells, and a subset of cells in the inner nuclear layer (bipolar cells), but not in horizontal cells (Brandst~itter et al., 1994). GluR7 transcripts are in all cell types except horizontal cells (Hamasssaki-Britto et al., 1993). KA2 mRNA is in all cell types (Brandst~itter et al., 1994). The KA1 gene is not expressed in the retina (Brandst~itter et al., 1994). Thus a major kainate receptor class in the retina might be GluR7/KA2 heteromeric assemblies (for example in bipolar cells, amacrine cells and ganglion cells), with GluR5/KA2 (for example in horizontal cells and ganglion cells), and GluR6/KA2 combinations occurring in other cell subsets (Fig. 10).

6. NEOCORTEX

Glutamatergic transmission in the neocortex is reviewed by Somogyi et al., 1998. The neocortex contains many cell types: glutamatergic pyramidal cells and many types of GABAergic interneuron (e.g. spiny stellate cells, basket cells and axo-axonic cells) which innervate the pyramidal cells and each other (Somogyi et al., 1998; Gupta et al., 2000). Double-labelling studies with glutamate receptor and marker probes (e.g. for Ca2+-binding proteins, neuropeptides or neuronal nitric oxide synthase) are essential in finding out which cell types express which glutamate receptor subtypes (e.g. Catania et al., 1995, 1998; Standaert et al., 1996, 1999). 6.1. NMDA RECEPTOR SUBUNIT mRNAs IN THE NEOCORTEX NR1 gene expression in the neocortex is strong (Figs. 5 and 7) (e.g. Moriyoshi et al., 1991; Monyer et al., 1992; Conti et al., 1994b; Laurie and Seeburg, 1994; Rudolf et al., 1996; Watanabe, 1997; Nase et al., 1999). In fact, NR1 subunit mRNA is in about 80% 113

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of rat cortical neurons (Conti et al., 1994b). The NR1 signal is highest in layers II and III and in V and VI. Layer IV has a lower signal. The NR1 splice variants (see Fig. 6) are differentially expressed in the cortex (Laurie and Seeburg, 1994; Laurie et al., 1995; Rudolf et al., 1996). The N-terminal splice (exon 5 insertion, NRI-b mRNA) has enriched expression in the somatosensory cortex (particularly in layer II, but also in V and VI) relative to other cortical areas; this somatosensory NRI-b expression is even more striking in the early weeks of postnatal development, especially for the outer cortical layers (Laurie and Seeburg, 1994). However, a large proportion of cortical NR1 mRNA (NRI-a) does not contain the exon 5 insertion (Fig. 7) (Laurie and Seeburg, 1994). For the C-terminal splices, the most abundant mRNA versions are those encoding both the C1 and C2 cassettes (NRI-1) and just the C2 cassette (NR1-2); a truncated version with neither C1 or C2 (NR1-4) is moderately expressed; the mRNA version with only the C1 cassette (NR1-3) is hardly detectable (Laurie and Seeburg, 1994; Laurie et al., 1995) (Fig. 7). Of the four NR2 subunit genes, only NR2A, NR2B and NR2D are significantly expressed in the rodent neocortex (Fig. 5) (Kutsuwada et al., 1992; Monyer et al., 1992, 1994; Rudolf et al., 1996; Watanabe, 1997; Nase et al., 1999; Standaert et al., 1999). NR2C probes give no detectable neocortical signal on X-ray film, but from emulsion studies the NR2C gene is weakly expressed in glial-like cells, i.e. small cells with scant cytoplasmic staining (Rudolf et al., 1996; Standaert et al., 1999). Both the NR2A and NR2B genes resemble NR1 in their expression: highest mRNA levels in layers II, III and VI (Fig. 5). There may, however, be slight differences between cortical areas. For example, in the prefrontal cortex, NR2A expression is highest in layers II-V, with II and III having a slightly stronger signal than V and Via; however, signal in VIb was much lower (Rudolf et al., 1996). The NR2B gene has highest expression in layers II and III and a less intense and more uniform expression in the deeper layers. Clearly, many NMDA receptor subtypes exist in the neocortex (Sheng et al., 1994): pyramidal cells are likely to have NR1/NR2A/NR2B-type receptors (i.e. high MgZ+-sensitivity to voltage, high channel conductance). The NR2A/NR2B subunit mRNA ratio increases during postnatal development (Fig. 8) (Monyer et al., 1994; Nase et al., 1999). In some neocortical areas, NMDA receptor gene expression is plastic, and so the NMDA receptor subunit composition can change in response to environmental stimuli (Nase et al., 1999). For example, in rat visual cortex, in situ hybridization experiments show that layer IV pyramidal cells regulate NR2A gene expression in proportion to the amount of sensory input they receive (Nase et al., 1999); this increase in the NR2A/NR2B ratio decreases the mean channel open time of the receptors. 6.2. NMDA RECEPTOR SUBUNIT mRNAs IN NEOCORTICAL INTERNEURONS By double-labelling ISH, somatostatin-, parvalbumin- and GAD67-positive cells express NR1, NR2A and NR2B, but not NR2C mRNA (Standaert et al., 1999). Unlike the other subunits, the NR2D gene seems to be expressed only in GABAergic interneurons. NR2D mRNA is in scattered cells in all laminae, which from double-labelling ISH are parvalbuminand somatostatin/NOS-containing GABAergic interneurons (Rudolf et al., 1996; Standaert et al., 1996, 1999); in contrast, neocortical enkephalin-positive interneurons may lack NR2D mRNA (Standaert et al., 1996). Some interneurons may have NR1/NR2D receptors (low Mg 2+ sensitivity, low channel conductance, long channel open time; Monyer et al., 1994), or receptors with more than one NR2 subunit. For these latter receptors one of the NR2 subunits may dominate the receptor properties. 114

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6.3. NR3A EXPRESSION IN NEOCORTEX The NR3A gene is expressed mainly in the deeper layers of adult neocortex with a punctate pattern (Fig. 9) (Ciabarra et al., 1995), suggesting strong expression in scattered cells. It is unknown whether these are pyramidal cell subtypes or GABAergic interneurons. This expression is stronger during the first postnatal weeks (Ciabarra et al., 1995). 6.4. AMPA RECEPTOR SUBUNIT mRNAs IN THE NEOCORTEX All four AMPA receptor subunit genes are expressed in the rat neocortex (Figs. 1, 2 and 11). GluR-D transcripts are rarer than the others (Kein~nen et al., 1990; Sato et al., 1993; Conti et al., 1994a; Gold et al., 1997). The expression patterns of GluR-A, -C and -D mRNAs differ among layers (Fig. 11) (Keinfinen et al., 1990; Sato et al., 1993; Conti et al., 1994a; Gold et al., 1996, 1997). By X-ray film autoradiography, GluR-D expression, as the flop splice version, is enriched in cortical layer IV (Figs. 2 and 11) (Keinfinen et al., 1990; Sommer et al., 1990; see fig. 3, plate L of Monyer et al., 1991). In Fig. 11, the GluR-D layer IV stripe is

Y

Fig. 11. AMPA receptor subunit mRNAs in the neocortex and caudate putamen of the adult rat (X-ray film, coronal sections). (A), GluR-A; (B), GluR-B; (C), GluR-C; (D), GluR-D. cc, corpus callosum; CPu, caudate putamen; ctx, neocortex; S, septum. Roman numeral indicates cortical layer. The arrows in D highlight the GluR-D expression in layer IV (Wisden and Seeburg, unpublished).

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arrowed. From double-labelling (GFAP immunocytochemistry combined with GluR-D ISH), neocortical GluR-D expression is in both astrocytes (cf. GluR-D expression in cerebellar Bergmann glia; Section 9.2.2) and neurons (Conti et al., 1994a). GluR-A and -C mRNAs are expressed strongly in layers II, III, V and VI, whereas the expression level of these genes in layer IV is lower; this is pronounced for GluR-C expression (Fig. 4) (Kein~nen et al., 1990; Sato et al., 1993; Conti et al., 1994a; Gold et al., 1996, 1997). Most cortical neurons have GluR-B mRNA, although as for the GluR-A and GluR-C genes, expression is lower in layer IV (Fig. 11) (Keinfinen et al., 1990; Sato et al., 1993; Conti et al., 1994a; Gold et al., 1996). For example, of 1426 cortical cells, 1139 were GluR-B mRNA-positive (Conti et al., 1994a). By quantifying silver grain intensity, GluR-B expression is highest in layer V, but is also strong in II, III and VI (Gold et al., 1996). Most GluR-B-expressing cells are pyramidal cells, and most of these do not contain, or have low levels of, GluR-A mRNA (Kondo et al., 1997). Kondo et al. performed a double-labelling study with digoxygenin-labelled cRNAs and AMPA receptor subunit-selective antibodies. For each cortical layer they selected randomly 200 cells. For GluR-A/GluR-B double-positive cells there were 10% in layers II and III, 8% in layer IV, 15% in layer V and 4.5% in VI; these GluR-A/GluR-B double-positive cells are both pyramidal and non-pyramidal cells (Kondo et al., 1997; for more on non-pyramidal cells, see Section 6.5). For GluR-A-negative/GluR-B-positive cells, there were 58% in layers II and III, 61.5% in layer IV, 55.5% in V and 62% in VI (Kondo et al., 1997). In layers II, III, V and VI these GluR-A-negative/GluR-B-positive cells are mostly pyramidal; in layer IV, they are both pyramidal and non-pyramidal (Kondo et al., 1997). These layer IV non-pyramidal cells may be a modified pyramidal cell type termed 'spiny stellate'. Flip splice versions of the GluR-A, -B, and -C mRNAs are distributed in a laminated pattern (Figs. 2 and 12), with highest expression in layers II, V and VI; flop expression is more uniform (Sommer et al., 1990; and figs. 2 and 3 of Monyer et al., 1991). By in situ hybridization, most of the cortical GluR-C expression is in the flip form (Sommer et al., 1990; Monyer et al., 1991). GluR-A flip, GluR-B flip and flop and GluR-C flip mRNAs are prominent in layer II (Figs. 2 and 12). According to a single-cell RT-PCR study, 90% of pyramidal cells in layers II and III use mostly flip variants, whereas 90% of layers II and III non-pyramidal cells (defined as fast spiking interneurons; see Section 6.5) use mostly flop variants (Lambolez et al., 1996). This matches the data shown in Fig. 12 for GluR-A flip and flop. 6.5. AMPA RECEPTOR SUBUNIT mRNAs IN NEOCORTICAL INTERNEURONS Certain GABAergic interneurons in the rat neocortex express mainly just the GluR-A flop and GluR-D flop subunits, with little or no GluR-B expression (Jonas et al., 1994; Catania et al., 1995; Geiger et al., 1995, 1999; Angulo et al., 1997; Kondo et al., 1997). Only 10-15% of cortical neurons are non-GluR-B- or low GluR-B-expressing cells (Kondo et al., 1997). For example, nitric oxide synthase (NOS)-positive neurons, identified with fluorescent secondary antibodies, contain only GluR-A and -D mRNAs as determined by hybridizing digoxygenin-labelled cRNA probes to the same sections (Catania et al., 1995). According to Kondo et al.'s double-labelling study, there are 9.5% GluR-A-positive/GluR-B-negative cells in layers II and III, 11.5% in layer IV, 13% in V and 13.5% in VI; most are non-pyramidal and express parvalbumin (Kondo et al., 1997). The 'no or little GluR-B' rule for GABAergic interneurons is not absolute. GluR-A/GluR-B double-positive cells in layers II-VI are mainly bipolar or multipolar and have intense calbindin-DzsK immunoreactivity (Kondo et al., 1997). Some of these cells may be the bipolar 116

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Fig. 12. The distribution of GluR-A flip and flop mRNAs in the adult rat neocortex (emulsion autoradiographs). (A), Nissl stain; (B), GluR-A flip; (C), GluR-A flop. Roman numerals indicate cortical layers. Arrows indicate examples of labelled cells. A strong band of flip-expressing cells is present in layer II of panel B (Wisden and Seeburg, unpublished).

GABAergic VIP-positive interneurons, which by single-cell RT-PCR contain mainly GluR-A flop and GluR-B flop m R N A s (Porter et al., 1998). A single-cell RT-PCR study on cortical cells defined as 'regular-spiking non-pyramidal' found mainly GluR-C flip and GluR-B flop expression (Angulo et al., 1997). 117

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ad Fig. 13. Expression of GluR5 mRNA in the developing neocortex (X-ray film autoradiographs). White arrowheads indicate the particularly intense line of expression in layer II cells in postnatal development. El7, embryonic day 17" P0, day of birth; P5, 5 days after birth; P12, 12 days after birth; ad, adult; CPu, caudate putamen. I, II and VI, neocortical layers. Scale bar, 0.7 mm (Bahn et al., 1994).

6.6. SUMMARY Most rat neocortical neurons have Ca2+-impermeable AMPA receptors containing GluR-A/B, B/C or A/B/C heteromeric assemblies, depending on cell type and the cortical layer; heteromeric receptors will be least numerous in layer IV cells (Fig. 11) (discussed by Conti et al., 1994a). Most GABAergic interneurons have CaZ+-permeable AMPA receptors made from GluR-A/D subunits; these receptors will have fast kinetics and high single-channel conductance. 6.7. KAINATE RECEPTOR SUBUNIT mRNAs IN THE NEOCORTEX Kainate receptor subunit gene expression in the neocortex is illustrated in Fig. 4. KA1 mRNA is also in the underlying corpus callosum white matter tracts (Werner et al., 1991; Wisden and Seeburg, 1993a; Bahn et al., 1994). By contrast, KA2 transcripts are abundant in the neocortex, particularly in layers II/III and V/VI (Herb et al., 1992; Wisden and Seeburg, 1993a; Bahn 118

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et al., 1994). The next most abundant neocortical kainate receptor subunit mRNA is GluR7. The confined expression of the GluR7 gene to the inner neocortical layers of all regions is striking (Fig. 4) (Bettler et al., 1992; Lomeli et al., 1992; Wisden and Seeburg, 1993a; Bahn et al., 1994). GluR7 mRNA in the deep layers is probably in pyramidal cells; in layers II and III there are also a few intensely labelled neurons (Wisden and Seeburg, 1993a). There are regional variations in GluR7 expression: certain regions show an intense but thin sublayer of GluR7 expression, possibly in layer III (see arrowheads in the GluR7 panels of Fig. 4) (Lomeli et al., 1992). GluR5 expression is present in scattered cells, possibly GABAergic intemeurons, in all cortical layers (except layer I). As for GluR7, GluR5 expression varies with cortical region (Fig. 13) (Bahn et al., 1994). In particular, the somatosensory cortex expresses more GluR5 transcript than other cortical areas, with highest levels in the outer layers (II and III) (Bahn et al., 1994). A peak of GluR5 expression in the somatosensory cortex is found around birth (Bahn et al., 1994). This expression is particularly high in layers II and III of the cortex: it correlates with the development of barrel fields (Fig. 13). Cortical GluR6 expression is uniformly weak; however, by single-cell RT-PCR, GluR6 mRNA, together with GluR5 mRNA, is found in a subset of GABAergic VIPergic intemeurons (Porter et al., 1998). There is a mismatch between the distribution of KA2 and the more limited distribution of the other subunits, suggesting unknown partner subunits for KA2 containing receptors in many parts of the cortex, particularly in the outer layers. The 31 subunit is weakly expressed in all layers of the cortex (Yamazaki et al., 1992); 82 mRNA is undetectable (Lomeli et al., 1993).

7. HIPPOCAMPUS Glutamate's importance in the hippocampus, the brain region essential for declarative memory formation, is emphasized by Bliss and Collingridge (1993). Like the cerebellum, the hippocampus is a region where a well-defined organization simplifies the description of receptor expression in the main cell types (pyramidal and dentate granule cells). There is only limited information, however, for glutamate receptor expression in GABAergic interneurons (Sommer et al., 1990; Monyer et al., 1991; Catania et al., 1995; Racca et al., 1996; Standaert et al., 1996, 1999). These GABAergic cells represent just 10% of hippocampal neurons, but they control the entire hippocampal network by feed-forward and feed-back inhibition (reviewed by Freund and Buzsaki, 1996). The many intemeuronal types (Freund and Buzsaki, 1996; Stevens, 1998; Geiger et al., 1999; Miles, 2000), located in the strata oriens, radiatum and lacunosum-moleculare, are impossible to identify from ISH alone, there are just too many subtypes. Double-labelling studies are needed (e.g. Catania et al., 1995, 1998; Standaert et al., 1996), but there are not yet enough cell-type-specific markers, and new cell types are regularly discovered. As for the GABAergic intemeurons in the neocortex and caudate putamen, broad categories can be defined by CaZ+-binding protein (e.g. parvalbumin, calretinin), peptide (somatostatin or NPY) or neuronal nitric oxide synthase expression. 7.1. HIPPOCAMPAL NMDA RECEPTORS

7.1.1. NMDA receptor gene expression in hippocampal principal cells The NR1, NR2A and NR2B genes are all highly expressed in adult dentate granule cells, and in the CA1-CA3 pyramidal cells (Monyer et al., 1992; Watanabe, 1997; Figs. 5, 8 and 119

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14); the NR2C and NR2D genes, however, are not expressed in these cell types (Monyer et al., 1994; Standaert et al., 1996). [In fact, the NR2C gene is expressed in small cells which may be glia (Fig. 14).] Pyramidal cells and dentate granule cells likely assemble NR1/NR2A, NR1/NR2B and/or NR1/NR2A/NR2B receptors. Most of the NR1 mRNA splice variants (see Fig. 6) are expressed uniformly in adult hippocampal pyramidal and dentate granule cells (Fig. 7) (Laurie and Seeburg, 1994; Laurie et al., 1995; Paupard et al., 1997). However, the N-terminal exon 5 insertion (NRI-b) has enriched expression in just CA3 pyramidal cells (Fig. 7), and the NR1-3 C-terminal version (only the C1 cassette present) is barely detectable (Fig. 7) (Laurie and Seeburg, 1994; Laurie et al., 1995; Paupard et al., 1997). From the X-ray film figure of Ciabarra et al. (1995), NR3A subunit mRNA is present in the CA1 pyramidal cell layer (Fig. 9).

7.1.2. NMDA receptor subunit gene expression in GABAergic interneurons By double-label ISH, somatostatin-, and parvalbumin-expressing cells in CA1, CA3 and hilus all contain NR1, NR2A and NR2B, but not NR2C mRNA (Standaert et al., 1999). Fig. 14 shows that there is little NR2A or NR2C signal in CA4 hilar cells, whereas the NR2B mRNA is in most cells in the CA4 area, including those under the dentate gyms blade; presumably some of this NR2B signal is due to interneuronal expression. As for the neocortex and caudate putamen, hippocampal NR2D subunit gene expression is confined to GABAergic cells. For example, in the CA1 and CA3 stratum oriens, many parvalbumin-, and somatostatin-positive cells contain NR2D mRNA (Standaert et al., 1996). NR1/NR2D receptors activate and deactivate slowly, during seconds rather than hundreds of milliseconds, and are less sensitive to voltage-dependent Mg 2+ block than NR1/NR2A or NR1/NR2B receptors (Monyer et al., 1994; Wyllie et al., 1998). Thus NR1/NR2D receptors may initiate action potentials on binding glutamate, even when the neuron is not substantially depolarized. However, hippocampal GABAergic basket cells, which by RT-PCR contain NR2B and NR2D mRNAs, do not have the NR2D-type response of long kinetics and weak Mg 2+ block (Catania et al., 1996). As hippocampal GABAergic cells express the NR2B subunit, this may dominate the properties of an NR1/NR2B/NR2D complex (cf. the NMDA response of cholinergic interneurons in the caudate putamen and other cells, which also express NR1, NR2B and NR2D mRNAs (Section 8.1.3) G6tz et al., 1997). 7.2. HIPPOCAMPAL AMPA RECEPTORS

7.2.1. AMPA receptor subunit gene expression in hippocampal principal cells The GluR-A, -B and -C genes express strongly in all hippocampal pyramidal cells and dentate granule cells (Boulter et al., 1990; Kein~inen et al., 1990; Pellegrini-Giampietro et al., 1991; Sato et al., 1993; Catania et al., 1995, 1998; Gold et al., 1996, 1997; Racca et al., 1996). GluR-D expression is confined to dentate granule cells and CA1 pyramidal cells (Bettler et al., 1990; Kein~inen et al., 1990; Sato et al., 1993; Catania et al., 1998).

+

Fig. 14. Expression of the NMDA receptor NR2 subunit mRNAs in the dentate gyrus of the adult rat hippocampus (emulsion autoradiographs). DG, dentate granule cells; arrows indicate examples of labelled cells. (Mower et al., 1992, 1994). Scale bar, 50 btm. 121

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7.2.1.1. Flip and flop RNA splicing in hippocampal principal cells The hippocampus has clear differences in the distribution of the AMPA receptor subunit flip and flop RNA splice variants (Figs. 2 and 15) (Sommer et al., 1990; Monyer et al., 1991). Predicting the subunit composition of AMPA receptors with respect to flip and flop isoforms is difficult. CA1 pyramidal cells express all flop versions (GluR-A to -D) and all flip versions, except GluR-D flip. GluR-C flop and GluR-D flop are relatively weakly expressed in CA1 cells. In contrast, CA3 pyramidal neurons synthesize only the flip version of GluR-A, GluR-B and GluR-C; flip mRNA levels are higher in CA3 than CA1 pyramidal cells (Sommer et al., 1990). No flop versions are detected in CA3 pyramidal cells by ISH. Dentate granule cells express all flip and flop forms with the exception of GluR-D flip, with flop mRNA levels higher than flip mRNAs (Fig. 2) (Sommer et al., 1990; Monyer et al., 1991). The ratio of flip to flop expression is different for GluR-A, -B and -C, e.g. GluR-C flip mRNA in dentate granule cells is more abundant than GluR-A flip mRNA (Kamphuis et al., 1994). Pyramidal and dentate granule cells might assemble different receptor configurations depending upon the subcellular location, e.g. dendrites versus soma (Wenthold et al., 1996; Yin et al., 1999).

7.2.1.2. Development of AMPA receptor flip and flop RNA splicing in hippocampal principal cells The flip and flop mRNA splice variants appear at different times during development. The flop versions are expressed at low levels prior to postnatal day 8; their characteristic high expression in CA1 pyramidal cells becomes apparent only during the second postnatal week (Fig. 16) (Monyer et al., 1991). In contrast, flip RNA levels are already substantial in pyramidal cells from birth (Monyer et al., 1991). As the CA1 pyramidal cells mature, the recruitment of the flop cassette into the AMPA receptors may cause these receptors to il)activate faster (Mosbacher et al., 1994). 7.2.2. AMPA receptor subunit mRNA in hippocampal interneurons Numerous interneurons in the hippocampus are labelled with GluR-A and -D probes (Figs. 15 and 17) (Sommer et al., 1990; Monyer et al., 1991; Catania et al., 1995, 1998), and there are some cells (especially in the stratum oriens) which are strong GluR-C expressors (Monyer et al., 1991; Catania et al., 1998). Many putative interneurons in the oriens, pyramidal and radiatum layers in both the CA1 and CA3 sectors strongly express subunits as flop, but not flip, variants (Fig. 15) (Monyer et al., 1991). Colocalization with glutamic acid decarboxylase-67 antibodies and digoxygenin-labelled GluR-B cRNA probes show that GluR-B mRNA is also in most GABAergic interneurons, but at lower levels than in pyramidal or dentate granule cells (Racca et al., 1996). This correlates with single-cell recording and RT-PCR studies on cultured hippocampal interneurons (Bochet et al., 1994) or in interneurons from slices (Geiger et al., 1995): GluR-A subunits dominate interneuron AMPA receptors, but they also contain GluR-B to -D flop subunit mRNAs. Catania et al. performed a double-labelling study to correlate AMPA receptor subunit gene expression with either parvalbumin or calretinin-expressing GABAergic interneurons (Catania et al., 1998). Parvalbumin-positive cells had high levels of GluR-A, GluR-C and GluR-D mRNAs, and low levels of GluR-B mRNAs; calretinin-containing spiny neurons express high GluR-A and GluR-D levels, low levels of GluR-B, and low/undetectable levels of GluR-C; calretinin aspiny neurons express low levels of GluR-A and GluR-D (Fig. 17) 122

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~~ ~ p y

Fig. 15. GluR-A flip and flop subunit mRNA expression in hippocampal interneurons in the CA3 and dentate gyrus areas of the adult rat hippocampus (emulsion autoradiographs). Left-hand column is dark field illumination; right-hand column is the corresponding Nissl stain under bright field. (A and B), GluR-A flop mRNA in the CA3 region; mRNA is absent from the pyramidal layer, but is detectable in certain non-pyramidal cells (interneurons) in the oriens and radiatum layers (arrows); (C and D), flip mRNA in CA3 pyramidal cells; (E and F), GluR-A flop mRNA is in dentate granule cells and in putative interneurons under the blade of the dentate gyrus (arrows). (G and H) GluR-A flip mRNA is weakly expressed in the dentate granule cells, and more abundantly in CA4 pyramidal cells. DG, dentate granule cells; Or, stratum oriens; Py, stratum pyramidale; Rad, stratum radiatum. Scale bar in H, 150 rtm (Monyer et al., 1991; Wisden, Seeburg and Monyer, unpublished).

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Fig. 16. The developmental expression of GluR-C flop in the rat hippocampus during the first two postnatal weeks (X-ray film, horizontal sections). Arrowheads indicate examples of labelled cells. Hi, hippocampus; Ctx, cortex; DG, dentate granule cells; Ent, entorhinal cortex; S, subiculum; PRh, perirhinal cortex. Scale bar, 0.8 mm (Monyer et al., 1991; Wisden, Seeburg and Monyer, unpublished).

(Catania et al., 1998). Similarly, from double-labelling with nitric oxide synthase antibodies and digoxygenin-labelled cRNA probes, most NOS-immunopositive hippocampal GABergic cells have high levels of GluR-A and -D, but not GluR-B or -C mRNAs (Catania et al., 1995). 124

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//

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Fig. 17. Drawings showing the relative expression levels of AMPA receptor subunit GluR-A to -D mRNAs in parvalbumin- (PV) and calretinin- (CR) immunopositive cells of the adult rat hippocampus (non-radioactive in situ hybridization combined with indirect fluorescence immunocytochemistry). Each dot represents one cell. Intensity levels are: black, strongly positive; grey, moderate or lightly labelled; white, negative (Catania et al., 1998).

This suggests that hippocampal GABAergic interneurons have highly Ca2+-permeable channels with fast kinetics. 7.3. KAINATE RECEPTORS AND 8 SUBUNIT IN THE HIPPOCAMPUS

7.3.1. Kainate receptor subunit mRNA expression in hippocampal principal cells KA1 mRNA is restricted to CA3 pyramidal cells and dentate granule cells (Fig. 4) (Werner et al., 1991; Wisden and Seeburg, 1993a; Bahn et al., 1994; Bureau et al., 1999). There is little KA 1 expression in CA 1 pyramidal cells. KA2 mRNA is abundant in both CA 1 and CA3 pyramidal cells and in the dentate granule cells. The GluR6 gene is moderately expressed in all CA pyramidal cells and in the dentate granule cells, with expression in CA3 higher than in CA1 (Egebjerg et al., 1991; Wisden and Seeburg, 1993a; Bureau et al., 1999; Paternain et al., 2000). GluR7 mRNA is in dentate granule cells but absent from CA pyramidal cells (Bettler et al., 1992; Lomeli et al., 1992; Wisden and Seeburg, 1993a). The 81 subunit gene is weakly expressed in CA1 and CA3 125

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pyramidal cells and dentate granule cells (Fig. 3) (Yamazaki et al., 1992; Lomeli et al., 1993). Possible kainate receptor subunit configurations in the principal cells are: GluR6/KA2 in CA1 pyramidals; GluR6/KA2 or GluR6/KA1 or GluR6/KA1/KA2 receptor(s) in CA3 pyramidals; receptors derived from KA1, KA2, GluR6 and GluR7 in dentate granule cells (Table 1).

7.3.2. Kainate receptor subunit mRNA expression in hippocampal interneurons There is little overall GluR5 mRNA expression in the rodent hippocampus seen by X-ray film autoradiography, but there is a strong punctate signal scattered in the subiculum, CA1 area (stratum oriens) and the dentate gyrus (Fig. 4) (Bettler et al., 1990; Wisden and Seeburg, 1993a; Bahn et al., 1994; Bureau et al., 1999; Paternain et al., 2000). This punctate pattern, especially clear during early postnatal development, is due to strong GluR5 expression in interneurons (Bahn et al., 1994; Bureau et al., 1999). This has been confirmed by double-labelling with GluR5 and GAD65 dixoxygenin-labelled cRNA probes (Paternain et al., 2000); approximately half of the interneurons (539 out 1004 cells evaluated) in adult CA1 stratum oriens express GluR5 (Paternain et al., 2000). This fits with the demonstration of functional GluR5 receptors on these cells (Cossart et al., 1998; Frerking et al., 1998). Some of these GluR5-positive cells are probably oriens-alveus-lacunosum-moleculare (OALM) interneurons. There are a few GluR5-positive cells in the stratum radiatum (approx. 14% of all GABAergic cells), and in the pyramidal cell layer itself (approx. 30% of all GABAergic cells located in the pyramidal cell layer; Paternain et al., 2000). According to Paternain et al, most of the GABAergic cells in the pyramidal cell layer also express GluR6, and so this specific subset of interneurons would be GluR5/GluR6-positive. There is a technical caveat: the spread of blue reaction product from the high GluR6 expression in pyramidal cells makes it difficult to be sure that interneurons in the pyramidal cell layer are really GluR6-positive (cf. Golgi cells in the cerebellar granule cell layer). There are a few GluR6-positive cells (6% of total GABAergic cells) in CA1 stratum oriens and stratum radiatum (approx. 3%); however, there are many GluR6-positive cells in CA3 stratum lucidum (Paternain et al., 2000); 85% of these GluR6-expressing stratum lucidum cells are also GAD65-positive. According to Bureau et al. there are a few GluR6 mRNA-positive cells in the mouse stratum oriens and radiatum of both CA1 and CA3 (Bureau et al., 1999). GluR7 is expressed in occasional cells in the pyramidal cell layer; it is unknown if these are pyramidal cell subsets or interneurons (see fig. 2F and G of Lomeli et al., 1992). There are a few GluR7 mRNA-positive cells in the mouse stratum oriens and stratum radiatum (Bureau et al., 1999).

8. CAUDATE PUTAMEN A simplified caudate putamen circuit is shown in Fig. 18. This circuitry contributes to movement regulation. Glutamatergic afferents from the cortex, thalamus, amygdala and substantia nigra innervate the principal GABAergic cells (medium spiny neurons) (Gerfen and Wilson, 1996; Wilson, 1998). The medium spiny cells co-release either enkephalin, or substance P with GABA. Those that synthesize enkephalin project to the globus pallidus; those that synthesize substance P project to the substantia nigra. Medium spiny cells make up over 90% of the caudate putamen cell population; so if the caudate is homogeneously positive on ISH X-ray film autoradiographs, this invariably reflects gene expression in the spiny projection cells. Information flow through the caudate is regulated by a sparse but important interneuronal population: the giant cholinergic cells, and various GABAergic interneuron 126

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Fig. 18. The cell types in the rat caudate putamen and their expression of the AMPA, NMDA and kainate receptor subunit mRNAs (circuit diagram adapted from Wilson, 1998). All cell types receive glutamatergic afferent input (open triangles). TH, tyrosine hydroxylase (dopaminergic terminals).

types (somatostatin and NOS-, parvalbumin-, and calretinin-containing cells; Kawaguchi et al., 1995). These cell types are non-overlapping. As for the medium spiny neurons, the interneurons (both cholinergic and GABAergic) are innervated by the glutamatergic fibres coming in from outside the caudate putamen (Fig. 18). 8.1. NMDA RECEPTOR SUBUNIT mRNA DISTRIBUTION IN THE CAUDATE PUTAMEN NR1 mRNA is present in most cells in the caudate putamen (Augood et al., 1994; Landwehrmeyer et al., 1995), although somatostatin/NOS-containing interneurons give weaker 127

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hybridization signals (Augood et al., 1994; Landwehrmeyer et al., 1995; Standaert et al., 1999). The medium spiny neurons are NRl-immunoreactive, as are the large- and mediumsized aspiny interneurons (Bernard and Bolam, 1998); nNOS-positive neurons also stain for NR1 immunoreactivity (Weiss et al., 1998).

8.1.1. NR1 splice variants From X-ray film analysis, NRI-a (no exon 5 insertion), NRI-1 (C1 plus C2) and NR1-2 (C2 only) are the main NR1 splice variants expressed in the caudate, and there is no exon5-containing transcript detectable (Laurie and Seeburg, 1994; Landwehrmeyer et al., 1995; see Fig. 6 for nomenclature). Projection neurons and interneurons contain different NR1 isoforms (Weiss et al., 1998; Ktippenbender et al., 1999): the C1 segment is confined to projection cells; by immunocytochemistry, the exon 5 insertion (NRI-b) in the N-terminus is found only in parvalbumin interneurons. Consistent with X-ray film autoradiography, a dual label in situ hybridization study detected no expression of exon-5-containing NR1 transcripts in enkephalin-positive spiny projection neurons, or in the cholinergic or somatostatin interneurons (Landwehrmeyer et al., 1995). From separate immunocytochemical and in situ hybridization studies, projection neurons contain NRI-1 (C1 and C2) and NR1-2 (C2 only); cholinergic cells contain mainly just NR1-2 (Landwehrmeyer et al., 1995; Weiss et al., 1998; Ktippenbender et al., 1999).

8.1.2. NR2 subunit expression By X-ray film autoradiography, the NR2A and NR2B genes are expressed in the caudate putamen, with 2B levels higher than 2A (Fig. 5; Monyer et al., 1992; Landwehrmeyer et al., 1995; Watanabe, 1997). The NR2A signal is higher in lateral and rostral caudate areas; the NR2B signal is homogeneous (Landwehrmeyer et al., 1995). From silver grain emulsions, NR2A is expressed in all caudate putamen cells and the regional differences in the NR2A signal on X-ray film are caused by variations in expression level; NR2B is highly expressed in all cell types. Striatal neurons do not express NR2C; however, a subpopulation of small cells with scant cytoplasm are labelled; similar NR2C-expressing cells are in the neocortex and hippocampus, these cells may be glia (Landwehrmeyer et al., 1995). NR2D mRNA is found in a small number of clearly labelled medium- and large-sized neurons; by ISH double-labelling, NR2D-expressing cells were identified as the somatostatin, parvalbumin and cholinergic interneurons (Standaert et al., 1994).

8.1.3. Summary Thus it is likely that medium spiny projection cells (both enkephalin and substance P types) express NR1/NR2A/NR2B receptors (Standaert et al., 1999); the exact composition may subtly vary with the location in the caudate, as NR2A expression varies regionally. The interneuron cell types, on the other hand, express mainly NR2B and NR2D (Fig. 18) (Standaert et al., 1994; Landwehrmeyer et al., 1995). Their receptors are likely to be either NR1/NR2D, NR1/NR2B and/or NR1/NR2B/NR2D forms. In spite of these differences in NMDA receptor subunit expression between principal and interneuronal cell types, the NMDA responses of spiny projection and cholinergic interneurons are remarkably similar (G6tz et al., 1997). A possible explanation is that the NR2B subunit is dominant; alternatively, 128

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the NR2D subtype may be targeted to peripheral dendrites, and is thus absent from nucleated patches (G6tz et al., 1997). 8.2. AMPA RECEPTOR SUBUNIT mRNA DISTRIBUTION IN THE CAUDATE PUTAMEN The GluR-A (flip and flop) and GluR-B (flip and flop) mRNAs are highly expressed in most caudate putamen cells, i.e. in the medium-sized spiny neurons (Figs. 11 and 18) (Sommer et al., 1990; Sato et al., 1993; Bren6 et al., 1998), and all medium spiny neurons have immunoreactivity for GluR-A and GluR-B/C (Bernard et al., 1997). The related medium spiny projection neurons in the nucleus accumbens also contain GluR-A and-B mRNAs (Lu et al., 1999). GluR-C (as flip form), and GluR-D mRNA are found in many caudate putamen cells, but at lower levels than GluR-A and GluR-B (Fig. 11) (Sommer et al., 1990). Consistent with the high expression of GluR-B in medium-sized spiny neurons, their AMPA receptors are almost impermeable to Ca 2+ (G6tz et al., 1997); these receptors deactivate and desensitize slowly, resembling AMPA receptor gating in hippocampal and neocortical pyramidal cells (G6tz et al., 1997). There are no in situ hybridization reports describing AMPA receptor expression in cholinergic cells; however, immunocytochemistry shows that they express the GluR-A and GluR-D, but not the GluR-B/C genes (Bernard et al., 1997). Consistent with this, AMPA receptors of cholinergic interneurons are highly Ca2+-permeable (G6tz et al., 1997). These cholinergic cells do not express the CaZ+-binding proteins usually associated with neurons with highly CaZ+-permeable AMPA receptors: parvalbumin, calbindin or calretinin are all missing. Cholinergic cell AMPA receptors deactivate and desensitize fast, comparable to AMPA gating in hippocampal and neocortical interneurons (G6tz et al., 1997). Parvalbumin-positive GABAergic neurons have GluR-A, GluR-B/C and GluR-D immunoreactivity (Bernard et al., 1997), but these have not been studied directly by in situ hybridization. It might be expected that NOS/somatostatin-positive interneurons have mainly GluR-A and GluR-D subunit mRNAs, as found in other brain regions, e.g. the hippocampus (Catania et al., 1995); however, these cells have little AMPA receptor mRNA (Catania et al., 1995), and do not stain with AMPA subunit antibodies (Bernard et al., 1997). 8.3. KAINATE RECEPTOR mRNA DISTRIBUTION IN THE CAUDATE PUTAMEN KA2 and GluR6 mRNAs occur at significant levels in virtually all medium-sized neurons in the caudate putamen, and are also in cholinergic neurons (Fig. 4) (Wisden and Seeburg, 1993a; Bischoff et al., 1997; Wullner et al., 1997; Chergui et al., 2000); thus, a main kainate receptor subtype on the medium spiny projection cells is likely to be KA2/GluR6. GluR7 mRNA is present in a subpopulation of medium-sized cells (approx. 60% of total cells in the caudate) (Lomeli et al., 1992; Wisden and Seeburg, 1993a; Wullner et al., 1997).

9. CEREBELLUM

Glutamate is a key neurotransmitter in the cerebellum: mossy fibres onto granule cells; parallel fibres onto Purkinje and stellate/basket cells, climbing fibres onto Purkinje cells, and also mossy fibre and climbing fibre inputs onto Golgi cells all use glutamate (Fig. 19) (Voogd 129

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Fig. 19. The cell types in the adult rat cerebellum and their expression of the AMPA, NMDA and kainate receptor subunit mRNAs [circuit diagram adapted from: Bahn and Wisden (1997); Cull-Candy et al. (1998)]. Excitatory terminals are open circles marked ' + ' . Inhibitory terminals are filled triangles marked ' - ' , and their cells are marked GAD (glutamic acid decarboxylase).

et al., 1996). The distribution of NMDA and non-NMDA receptor subunit mRNAs in the cerebellum is summarized in Fig. 19. 9.1. NMDA RECEPTOR SUBUNIT mRNAs IN THE CEREBELLUM

9.1.1. Purkinje cells Adult Purkinje cells strongly express the NR1 gene (Watanabe et al., 1994; Cull-Candy et al., 1998). All the NR1 RNA splice variants are found, although NR1-3 (C1 only, see Fig. 6) RNA is at low levels (Laurie et al., 1995). However, according to most reports, adult Purkinje cells do not contain NR2-type subunit mRNA (Watanabe et al., 1994), and indeed have no detectable NMDA receptors (Cull-Candy et al., 1998); nevertheless, two groups have described NR2A mRNA in Purkinje cells (Akazawa et al., 1994; Luque and Richards, 1995). This has still not been resolved. The mismatch in expression between NR1 and the NR2 series has been much commented on (Cull-Candy et al., 1998), and it is indeed unusual to find a neuron type with no NMDA response. An outside possibility is that the NR1 subunit 130

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associates with the ~2 orphan subunit (see Section 9.3. l) to form a glutamate receptor channel with novel properties. The NR2D gene is expressed in young postnatal Purkinje cells along with NR1 (Akazawa et al., 1994) and between P 1 and P 10, low-conductance NMDA receptors are found (Cull-Candy et al., 1998). The 'juvenile' NMDA receptor expression on Purkinje cells can persist in some mouse mutants. In adult staggerer mice, NR2A, NR2D and NR1 mRNAs are all found in Purkinje cells, and this expression varies with the cerebellar domain (Nakagawa et al., 1996); the NR2-expressing Purkinje cells are separated from each other by narrow regions in which Purkinje cells have no NR2 subunit mRNAs; wild-type mouse Purkinje cells have only NR1 mRNA (Nakagawa et al., 1996). In the adult reeler mouse cerebellum, NR2A mRNA is found in those Purkinje cells located in the rostral zones (Watanabe et al., 1995); NR1 mRNA is found in all reeler Purkinje cells. These two mouse lines show that there is variability in Purkinje cells regarding expression of NMDA receptor subunits.

9.1.2. Bergmann glial cells NMDA-activated currents are found in Bergmann glia (Mfiller et al., 1993); rat Bergman glia cells have small amounts of NR2B mRNA, and also express the NR1 subunit mRNA (Luque and Richards, 1995). NR2A/B protein can be detected on Western blots of cultured chick Bergmann glia extract (Lopez et al., 1997).

9.1.3. Granule cells Adult granule cells transcribe the NR1, NR2A and NR2C genes. In fact, granule cells are the highest NR2C gene expression sites in the brain (Monyer et al., 1992; Akazawa et al., 1994; Watanabe et al., 1994). All the NR1 splice variants are present, but the NR1-3 mRNA signal is weak (Laurie et al., 1995). There is a switch in NMDA receptor subunit gene expression in developing postnatal granule cells. Young pre-migratory and post-migratory granule cells express the NR1 and NR2B genes, but during the second postnatal week, the NR2A and NR2C genes are turned on, and NR2B RNA levels decline (Monyer et al., 1994). The recruitment of NR2A into more mature NMDA receptors is found in many other brain regions as well, for instance in pyramidal cells of layer IV visual cortex (Monyer et al., 1994; Nase et al., 1999) (Fig. 8).

9.1.4. GABAergic interneurons Stellate and basket cells transcribe the NR1, and NR2D genes (Watanabe et al., 1994). This fits with the common theme of NR2D expression in GABAergic interneurons in other brain regions (cf. Sections 6.2, 7.1.2 and 8.1). The main NR1 splice variants are NRI-a and NRI-b, and NR1-4 (Laurie et al., 1995). From ISH, NMDA subunit gene expression in Golgi cells is not clear: the high labelling of the surrounding granule cells obtained with NR1, NR2A and NR2C probes interferes with seeing if the rare Golgi cells, whose cell bodies are scattered in the granule cell layer, are labelled. However, adult Golgi cells do not express NR2B; if they did this would be clearly seen, as adult granule cells do not express NR2B. An educated guess is that Golgi cells express NR1 and NR2D (Cull-Candy et al., 1998).

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9.1.5. Cerebellar nuclei The cerebellar nuclei express mainly the NR1, NR2A and NR2D subunit genes (Watanabe et al., 1994). The NR3A gene is expressed at low levels in the adult cerebellum (cell types unknown); by RT-PCR, the NR3-L splice version predominates (Sun et al., 1998). 9.2. AMPA RECEPTOR SUBUNIT mRNAs IN THE CEREBELLUM 9.2.1. Purkinje cells Purkinje cells express GluR-A flop, GluR-B flip and flop, and GluR-C flip mRNAs; GluR-A expression is the weakest (Kein~inen et al., 1990; Sommer et al., 1990; Monyer et al., 1991; Sato et al., 1993; see Fig. II-L for pan GluR-A to -D expression). There are thus probably multiple AMPA receptors on Purkinje cells: for example, these could be differentially located at parallel fibre and climbing fibre synapses. 9.2.2. Bergmann glial cells Bergmann glial cells express the GluR-A flip and -D flip mRNAs (Fig. 1) (Kein~inen et al., 1990; Sommer et al., 1990; Monyer et al., 1991; Burnashev et al., 1992; Gallo et al., 1992; Sato et al., 1993; Kondo et al., 1997). Bergmann glia have two types of GluR-D flip subunit mRNAs, differing by alternative splicing in the region encoding the C-terminus (Gallo et al., 1992). No functional differences have been demonstrated for these (Gallo et al., 1992). The GluR-B gene is not expressed in Bergmann glial cells, and so these cells assemble CaZ+-permeable AMPA receptors with fast kinetics and high single-channel conductance (Burnashev et al., 1992; MUller et al., 1992b; Geiger et al., 1995). 9.2.3. Granule cells Granule cells contain only GluR-B flip and GluR-D flop mRNAs (KeinS.nen et al., 1990; Sommer et al., 1990; Monyer et al., 1991; Sato et al., 1993). As for Bergmann glia, both C-terminal splice variants of the GluR-D gene combine with the flop module (Gallo et al., 1992). As granule cells mature, there is a switch of GluR-D transcript splicing. In rats younger than two weeks little GluR-D flop mRNA is detected by ISH, whereas GluR-D flip mRNA is prominent in the granule cell layer (Mosbacher et al., 1994). By the third week, there are higher levels of GluR-D flop, and GluR-D flip mRNA levels decline, with the electrophysiological properties of the receptors (faster desensitization) changing accordingly (Mosbacher et al., 1994). 9.2.4. GABAergic interneurons Stellate/basket cells contain GluR-B and GluR-C mRNAs; Golgi cells possibly have GluR-C mRNA (Kein~inen et al., 1990). Because of the high silver grain density obtained over the granule cells when using GluR-B and -D probes, it is not possible to see i f - B and -D transcripts are in the Golgi cells (Kein~inen et al., 1990; Sato et al., 1993).

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9.2.5. Cerebellar nuclei (medial, interposed and lateral) These contain GluR-B, GluR-C and GluR-D transcripts (Sato et al., 1993). 9.3. KAINATE RECEPTOR AND 8 SUBUNIT mRNAs IN THE CEREBELLUM

9.3.1. Purkinje cells KA1 and GluR5 are the only kainate receptor subunit mRNAs in Purkinje cells (Bettler et al., 1990; Werner et al., 1991; Wisden and Seeburg, 1993a; Bahn et al., 1994; Niedzielski and Wenthold, 1995). Thus, Purkinje cells might assemble KA1/GluR5 kainate receptors. Both mice and rat Purkinje cells have high levels of ~2 mRNA (Fig. 3) (Araki et al., 1993; Lomeli et al., 1993) and 8 immunoreactivity (Araki et al., 1993; Mayat et al., 1995). The 32 subunit, which is specifically at Purkinje cell dendritic spine/parallel fibre synapses (Zuo et al., 1997), contributes to the regulation/induction of long-term depression at the parallel fibre synapse; mice lacking this subunit are ataxic (Hirano et al., 1995; Kashiwabuchi et al., 1995). In the lurcher mouse, a dominant negative mutation in the 8 subunit produces excitotoxic death of Purkinje cells (Zuo et al., 1997). So based on both gene knockout and gain-of-function studies, the 82 subunit contributes to an ionotropic glutamate receptor. 82 may contribute to receptors with AMPA or kainate subunits, or possibly even assemble with the NR1 subunit.

9.3.2. Granule cells Granule cells express the KA2 and GluR6 subunit mRNAs (Egebjerg et al., 1991; Herb et al., 1992; Wisden and Seeburg, 1993a; Bahn et al., 1994; Niedzielski and Wenthold, 1995). Thus granule cells might assemble KA2/GluR6 receptors (Fig. 19). Some KA2 immunoreactivity is located on parallel fibres, suggesting that granule cell GluR6/KA2 receptors might function pre-synaptically (Petralia et al., 1994).

9.3.3. GABAergic interneurons Basket/stellate cells express moderate amounts of GluR7 RNA, but no other kainate subunit mRNAs (Lomeli et al., 1992; Wisden and Seeburg, 1993a). These cells probably assemble homomeric GluR7 receptors; on recombinant homomeric GluR7 receptors, glutamate has a 10-fold lower potency in producing currents compared with other non-NMDA receptor channels (Schiffer et al., 1997). Only high glutamate concentrations activate homomeric GluR7 channels: 1 mM glutamate causes tiny currents; 30 mM glutamate is needed for maximal currents (Schiffer et al., 1997). Even if basket/stellate cell GluR7 receptors are synaptic, it is unclear if glutamate concentrations reach high enough levels to activate them (Schiffer et al., 1997). The other 'GluR7 mismatch' occurs in the reticular thalamic nuclei (Lomeli et al., 1992).

10. SPINAL CORD Non-NMDA and NMDA receptors are used by many spinal neuronal cell types (reviewed by: Zieglg~insberger and T611e, 1993; Lodge and Bond, 1994; Woolf and Costigan, 1999). Ionotropic glutamate receptors are also expressed by nociceptive primary afferent neurons, i.e. 133

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Fig. 20. The distribution of the NMDA receptor subunit NR1 mRNA splice variants in the adult rat spinal cord (X-ray film autoradiographs, coronal sections). Scale bar, 300 gm (T611e et al., 1995a). See Fig. 6 for explanation of the nomenclature.

peripheral nerve cells whose soma are located in the dorsal root ganglia (Woolf and Costigan, 1999). Embryonic dorsal root ganglia (DRGs) have high levels of GluR5 and KA2 mRNAs and modest GluR7 mRNA levels (Bettler et al., 1990; Herb et al., 1992; Lomeli et al., 1992). GluR5 mRNA is also in adult DRGs (Bettler et al., 1990). The kainate receptors are on the DRG pre-synaptic terminals in the dorsal horn, where they control neurotransmitter release (Woolf and Costigan, 1999). 10.1. NMDA RECEPTOR SUBUNIT mRNAs IN THE LUMBAR SPINAL CORD High levels of NR1 mRNAs are found throughout the cord (Fig. 20) (T611e et al., 1993; Luque et al., 1994; Watanabe et al., 1994; Shibata et al., 1999; see Fig. 6 for splice variant 134

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nomenclature). By X-ray film autoradiography, the main NR1 RNAs are: NRl-a (no exon 5 insertion), NRI-b (exon 5 insertion), NRI-1 (C1 and C2) only in dorsal horn cells (laminae I to III), NR1-2 (C2 only) and NR1-4 (no C1 or C2); there is little NR1-3 (C1 only) mRNA (Luque et al., 1994; T611e et al., 1995a,b). NRI-b (exon 5 insertion) RNA is found mainly in dorsal horn neurons in laminae II and III, but also in some neurons in layers I and II-outer, and in some laminae IV and V cells (T611e et al., 1995a) (Fig. 20). Ventral horn motor neurons have mainly NRI-a, NR1-2 and NR1-4 RNAs. NRI-1 RNA, which gives no ventral horn signal on X-ray film autoradiography, is enriched in the nuclei of motor neurons; this may reflect part of the splicing control mechanism, and that this RNA is not available for translation (T611e et al., 1995a,b). Which NR2 mRNAs are found in the rat lumbar spinal cord? T611e et al. (1993) described only NR1 and NR2D mRNAs; however, three other groups found NR2A and NR2B mRNA as well, and so this is most likely correct (Luque et al., 1994; Watanabe et al., 1994; Shibata et al., 1999). There are moderate levels of NR2A mRNA in spinal grey matter, except for lamina II, and low but significant levels of NR2B mRNA in lamina II and some ventral horn motor neurons. NR2D mRNA is found throughout the grey matter at low levels (T611e et al., 1993; Luque et al., 1994; Watanabe et al., 1994; Shibata et al., 1999). NR2C probes gave faint signals in small cells dispersed over the grey and white matter, suggesting glial expression (Shibata et al., 1999). NR3A mRNA is present throughout the grey matter of both cervical and lumbar regions; the highest levels are in dorsal horn laminae II and III (Ciabarra et al., 1995); the strength of the NR3 signal has not been compared directly with the NR2 signals. In summary, many spinal cord neurons will use NR1/NR2A receptors, and there will be a minority of NR1/NR2B or NR1/NR2A/NR2B receptors. There is substantial variation in the NR1 splice forms used, and it is likely that many spinal cord neurons use multiple NR1 types. Unlike ventral horn motor neurons, visceromotor neurons express only the NR1 mRNA (cf. cerebellar Purkinje cells and retinal horizontal cells; Shibata et al., 1999). 10.2. AMPA RECEPTOR SUBUNIT mRNAs IN THE LUMBAR SPINAL CORD 10.2.1. Dorsal horn

GluR-A and GluR-B transcripts dominate in the dorsal horn; GluR-C and GluR-D dominate in the ventral horn (Fig. 21) (Furuyam et al., 1993; Sato et al., 1993; T611e et al., 1993, 1995b). GluR-A expression, mainly as the GluR-A flop splice type, is confined to dorsal horn laminae I and II-outer (T611e et al., 1993, 1995b; Tachibana et al., 1994). Many dorsal horn AMPA receptors are likely to contain GluR-B flip, possibly as GluR-B flip/GluR-A flop heteromerics in laminae I and II-outer, and GluR-B flip homomerics in laminae I and II with minor populations of C- and D-containing receptors (T611e et al., 1995b). Based on the prevalence of the GluR-B subunit mRNA and protein in the dorsal horn (Fig. 21), many AMPA receptors there are likely to be CaZ+-impermeable (Furuyam et al., 1993; T611e et al., 1993, 1995b; Tachibana et al., 1994). However, an AMPA receptor subpopulation on dorsal horn neurons in laminae I and II-outer is strongly CaZ+-permeable (Gu et al., 1996; Engelman et al., 1999). These cells are both GABAergic (interneurons) and non-GABAergic (NKl-receptor-expressing projection cells) and probably receive nociceptive input directly by glutamate from dorsal root ganglion cells (Albuquerque et al., 1999). It is unclear if these CaZ+-permeable receptors are GluR-A flip homomerics, or heteromers with GluR-A flip and fewer subunits of GluR-B (subunit composition is variable in AMPA receptors, and depends on the relative expression level of the subunits; Monyer et al., 1999). 135

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Fig. 21. (a-b) The expression of the AMPA receptor subunit genes (GluR-A, GluR-B, GluR-C and GluR-D) in the

adult rat lumbar spinal cord (X-ray film autoradiographs, coronal sections). Scale bar, 300 I~m. (T611eet al., 1993).

10.2.2. Ventral h o r n m o t o r n e u r o n s

The main AMPA receptor subunit mRNAs in the ventral horn are GluR-C and -D (Fig. 21) (Furuyam et al., 1993; Sato et al., 1993; T611e et al., 1993; Shibata et al., 1999). From serial sectioning and hybridizing consecutive sections, somatomotor neurons express GluR-B flip, GluR-C flip, GluR-C flop and GluR-D flip subunits, but GluR-B transcripts are less abundant than the GluR-C and -D mRNAs (T611e et al., 1993, 1995b) GluR-B transcripts are enriched in the cell nucleus (T611e et al., 1993). The retention of GluR-B transcripts in the nucleus may be related to the RNA editing process; however, GluR-B Q / R site editing is essentially 100% in motor neurons (Vandenberghe et al., 2000); alternatively, nuclear retention may help regulate translational availability of GluR-B mRNA. If GluR-B mRNA is present in limiting amounts in the cytoplasm, then motor neuron AMPA receptors might have moderate Ca 2+ permeabilities (Bochet et al., 1994; Jonas et al., 1994; Monyer et al., 1999). The presence of GluR-D flip rather than flop in motor neurons is unusual; the only other locality where GluR-D flip is abundant is cerebellar Bergmann glia (see above). These results agree with immunocytochemistry studies of ventral horn motor neurons, which have strong staining with GluR-D and with GluR-B/-C antibodies (Tachibana et al., 1994). Given the number, of subunit 136

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m R N A s (including splice forms) found in motor neurons, these cells probably assemble multiple A M P A receptor subtypes, some of which are CaZ+-permeable. This has been directly confirmed (Greig et al., 2000; Vandenberghe et al., 2000). Visceromotor neurons in the rat lumbosacral (L6-S 1) spinal cord express different A M P A receptor subunit m R N A s from ventral horn motor neurons, namely GIuR-A and -B, with little or no GluR-C or -D m R N A s (Shibata et al., 1999). Visceromotor neurons also differ from ventral horn motor neurons in their N M D A receptor subunit gene expression (Shibata et al., 1999; see above). 10.3. K A I N A T E A N D 3 R E C E P T O R S U B U N I T m R N A s IN T H E S P I N A L C O R D Kainate receptor subunit m R N A s are not abundant in the adult spinal cord, and GluR6 is not expressed at all (T611e et al., 1993). In the dorsal horn, occasional cells express the GluR5 and GluR7 subunit genes, and more cells contain KA2 m R N A (T611e et al., 1993). Kainate receptors are probably in subsets of A M P A receptor-positive cells. Most of the GluR5 protein in the dorsal horn is on the primary afferent terminals of D R G cells (Woolf and Costigan, 1999). Motor neurons express the KA1 gene, and weakly express the GluR5 gene (T611e et al., 1993). Both 3 subunit genes are weakly expressed throughout the cord's grey matter (T611e et al., 1993).

11. ACKNOWLEDGEMENTS We thank M a r y - A n n Starkey for help in preparing the manuscript.

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CHAPTER V

Regional and synaptic expression of ionotropic glutamate receptors R.S. PETRALIA, M.E. RUB IO, Y.-X. WANG AND R.J. WENTHOLD

1. INTRODUCTION Ionotropic glutamate receptors are made up of complexes of four or five subunits forming a central ion channel that passes sodium or calcium ions. They include the AMPA receptors, with four subunits, GluR1-4, the kainate receptors with five subunits, GluR5-7 and KA1-2, the delta receptors with two subunits, delta 1-2 (~ 1-2), and the NMDA receptors with six subunits, NR1 (~ 1), NR2A-D (~ 1-4), and NR3 (X-1 or NMDAR-L). Many subunits also have variant forms generated through alternative splicing. Different subunits within each group usually combine to form heteromeric receptor complexes, although homomeric complexes made entirely of one kind of subunit do occur. Thus, fully functional NMDA receptors require NR1 plus at least one kind of the NR2 subunits, while AMPA receptors can be heteromeric or homomeric complexes; the best example of the latter is GluR1 in certain neuron populations in several regions of the brain. This chapter begins with a survey of ionotropic glutamate receptor distribution in the brain and other organs (Hollmann and Heinemann, 1994; Petralia and Wenthold, 1996; Bahn and Wisden, 1997; Watanabe, 1997), and then continues with a discussion of the factors affecting expression of ionotropic glutamate receptors in synapses, mainly in the postsynaptic spine (Ottersen and Landsend, 1997; Petralia, 1997; Somogyi et al., 1998; Petralia et al., 1999b,c,d).

2. REGIONAL DISTRIBUTION This review will be limited to major structures in adult mammals; due to lack of space, invertebrates, lower vertebrates, and developmental stages (see Section 3.1.2) will be mentioned only incidentally. In the brain, discussion is limited to major nuclei; information on other nuclei can be found in Table 1 and in many of the earlier papers mentioned here. Also, due to the large number of publications on glutamate receptor distribution, coverage is limited mainly to the earlier comprehensive papers and representative work from the recent literature. In this section, most references are designated IS for in situ hybridization and IC for immunocytochemistry. Most studies used rats, although some used mice (especially the Watanabe et al. papers) or other mammals; animal species is mentioned when it may be important. Discussions of the levels of mRNA or protein expressed (low, moderate, high) typically are based on the amount relative to the highest labeling seen in each study; thus the results are not necessarily comparable between different receptor subunits. Handbook of Chemical Neuroanatom 3, Vol. 18: Glutamate O.P. Ottersen and J. Storm-Mathisen, editors Published by Elsevier Science B.V.

145

C~

TABLE 1. Expression of mRNA for glutamate receptors in a selection of CNS structures

Olfactory bulb (main) Glomerular layer Mitral layer Granular layer Anterior olfactory nucleus Piriform cortex (layer II*) Neocortex - laminae II/VI Hippocampus CA1 CA3 Dentate gyrus (granule cells*) Striatum Caudate putamen Globus pallidus Septum Bed n. stria terminalis (ven.*) Lateral septum (dorsal*) Medial septum Diagonal band Amygdala Lateral n. Basolateral n. Central nucleus Medial habenula Thalamus Reticular nucleus Ventral postero(medial*) n. Anteroventral n. Dorsolateral geniculate n. Hypothalamus Medial preoptic n. Dorsomedial n. Ventromedial n. Suprachiasmatic n.

NMDA receptor subunits

Kainate receptor subunits

AMPA receptor subunits R1

R2

R3

R4

R5

R6

R7

+++ +++ +++ +++ +++*

+++ +++ +++ +++ +++*

+ +++ +++ +++ +++*

+++ +++ +++ ++ +++*

+++

+++

+

+++

+++

+++

+/++

+

+

+/+++

+++

+++

+++

+++

+

+++ +++*

+++ +++*

++ +++*

+ 0 0

+

+++ +++*

+ ++

+ ++

KA1

KA2

NR1

N2A

N2B

N2C

++ +++

+ +

+ ++

+ +

+++ +++

+ +++

++ +++

0

N2D

+ +

+++ +++

+++ +++

+++ +++

+++ +++

0 0 0

+ +++ +++

+++ +++ +++

+++ +++ +++

+++ +++ +++

+++ +++ +++

0 0 0

0 0 0

+++

+++

++

++

+

+

++

++

+

+

++

+

0

+

+ +

+++ +

+++ +

+ +

++ +

0 0

0 +

+++ +++ ++ ++

+++ +++ ++ ++

++ +++ +++ +++

+ + ++ ++

+++ ++ +

+ + +

+ ++ ++

+ + +

+++ ++ ++

++* ++* ++ ++

O* +* + +

+* ++* + +

O* O* 0 0

O* O* + +

+++ +++ +++

+++ +++ +++

+++ +++ +

+ + +

+++

+++

+++

+++

0

0

+

+

+++

+++ +++ +++ +

++ +++ + +

+++ +++ + 0

0 0 0 0

0 0 0 0

0 + ++ ++

++ + ++ +

+++ +++ +/++ ++

+++ +++ ++ ++

0 0 +++

+

+++ 0 ++

+ 0 +

+ 0 ++

++

+

+

+

+

0

+++* +++ +++

+++ + ++

+++ ++ +++

+ + +

0 + +

+++

+++

+

+

+

+

+

+++ +++ ++

+++ +++ +++

+ +

+ +

+ +

0 0

+ ++

0

+

+++

0

+

+ + +++ +

+ 0 + 0

+ 0 + 0

0 0 0 +++

+ 0 0 0

0

+ + + +

++ ++ + ++

.ce t...,

~.~~

TABLE 1 (continued) o~

R1

Mammillary (med./lat.) n. Arcuate n. Substantia nigra Pars compacta Pars reticulata Red (parvocell./magnocell.) n. Superior colliculus Superficial gray layer Deep gray layer Inferior colliculus (central n.*) Oculomotor n. Trigeminal mesencephalic n. Trigeminal principal n. Motor trigeminal n. Facial n. Ventral cochlear n. (anterior*) Dorsal cochlear n. Superficial layer (layer 2*) Deep layers Vestibular nuclei Medial n. Other n. Nucleus of the solitary tract Hypoglossal n. (oral part*) Ambiguus n. External cuneate n. Cuneate n. Pontine n. Inferior olive (medial n.*) Dorsal raphe n. Pontine reticular n. Gigantocellular reticular n. Locus coeruleus -...3

R2

NMDA receptor subunits

Kainate receptor subunits

AMPA receptor subunits R3

R4

R5

R6

R7

KA1

KA2

NR1

N2A

N2B

N2C

N2D

+ +

+ 0

0 0

0 0

0 0

++ + +++

+ + +

+ 0 0

0 0 0

0 0 0

0/+++ +++

+++/+ +++

0 0

+ 0

++ ++ 0/++

+++ ++ +

+ + +++

++ ++ ++

+++ ++ + 0 0 + 0 +/++ 0

++ ++ +++ +++ +++ +++ +++ +++ ++

+ + +++ +++ ++ + +++ +++ +++

+ + + ++ +4+ +++ +++ +

+++ +++ +++* ++ ++ ++ +++ +++ ++*

+ + ++* + + + + + +*

+ 0 0* 0 0 0 0 0 0*

0 0 0* 0 0 0 0 0 0*

+ + 0* 0 0 0 0 0 0*

+++ 0

+++ +++

+++ +++

+ +

+++* +++

+* +

++* 0

++* 0

0* 0

+ 0/++ ++ +* + +++ ++

+++ ++ +++ +++* +++ ++ ++ ++ ++ +++ ++ ++ +++

+++ +++ + ++* ++ ++ + +++ + + ++ +++ ++

++ ++ + +++* ++ ++ ++ ++ ++ ++ ++ ++ +

0

+++ ++ ++ ++ +

0 0

0 +

+++ 0

0 0

++ +

+++

0

++

+

+

0

+

+

+

++

0

+

0

+

+++

+++

+

0

+

+

+/++

+

o

o

o/+

+++ +++ ++ +++ ++ +++ ++* ++ +++ +++ ++

+ ++ + + + + +++* + + + +

+ + 0 0 0 + +* 0 0 0 +

0 0 0 0 0 0 0* 0 0 0 0

0 0 0 0 0 0 0* + 0 0 0

t,,,~ ~

e,,~~

%

TABLE 1 (continued) Kainate receptor subunits

AMPA receptor subunits

Cerebellum Purkinje cells (layer*) Granule cells (layer*) Cerebellar nuclei Spinal cord Dorsal horn - laminae I-III Ventral horn - motoneurons

R1

R2

R3

R4

§ 0 0

§247247 §247247 § §247247 0 §247 §247247 §247247 §247

R5

R6

§247 0

§247247 §247247 §247 §247 § § §247247 §247247 §247247 §

R7

NMDA receptor subunits KA1

KA2

0* 0* §247247 0

§ 0

0 0

§ §247

§ 0

NR1

N2A

N2B

N2C

N2D

0* §247247 §247247 §247247 §247247

0* § §247

0* 0* 0

0* §247247 0

0* 0* §

§247 0

0 0

0 0

§ 0

+§ §247

§247247 §247247

This table is simplified from tables of AMPA (adult rat; Sato et al., 1993a), kainate (adult rat; Wisden and Seeburg, 1993) and NMDA (mouse, mainly P21; Watanabe, 1997) receptor subunits. All spinal cord results (adult rat lumbar segments) are from T611e et al. (1993); Watanabe (1997) reports somewhat different results for the mouse cervical segments (see text). Levels of kainate receptors in the substantia nigra (adult rat) are taken from Wtillner et al. (1997) - - also their values for the caudate putamen and globus pallidus are somewhat different from those of Wisden and Seeburg (1993). Classification of levels of mRNA expression have been simplified to '0' for no mRNA expressed, ' + ' for very low to low levels, ' + + ' for moderate levels, and ' § 2 4 7 2 4for 7 high to very high levels. For the Watanabe (1997) study, levels 1-2 are represented here by ' + ' , level 3 by ' + + ' , and levels 4-10 by ' § 2 4 7 in this case, levels were based on signal-to-noise ratios calculated for silver grain densities over unit areas. For the T611e et al. (1993) study, the rating used here is ' + ' for weakly detectable, ' § for detectable, and ' § 2 4 7 2 4for 7 abundant.

Regional and synaptic expression of ionotropic glutamate receptors

Ch. V

2.1. FOREBRAIN The olfactory bulb has moderate to high levels of GluR1-4 (IC, Petralia and Wenthold, 1992; IS, Sato et al., 1993a) and GluR5-7 (IS, Hollmann and Heinemann, 1994); low to moderate levels of KA2 also are present (IC, Petralia et al., 1994c). Moderate immunolabeling for delta 1/2 is seen, with highest levels in mitral cells (IC, Mayat et al., 1995). There are moderate levels of NR1, NR2A and NR2B, but only low levels of NR2C and NR2D (IS, P21, Watanabe et al., 1993; IC, Petralia et al., 1994a,b; Wenzel et al., 1996). Of the NR1 splice variants, those that lack the N1 cassette (= N1 segment) are common throughout, while those containing the N 1 cassette are most prevalent in the granule cells (IS, Laurie et al., 1995). Based on RT-PCR, the olfactory bulb may contain similar levels of short and long variants of NR3 (Sun et al., 1998). Also, substantial levels of NR3 are found in the nucleus of the lateral olfactory tract (IS, Sucher et al., 1995). In the neocortex, GluR1-3 are high while GluR4 is low (IC, Petralia and Wenthold, 1992; IS, Sato et al., 1993a) (Fig. 1). Many nonpyramidal neurons have GluR1 but little or no GluR2, which is responsible for calcium impermeability of AMPA receptor channels; this indicates that these neurons have calcium-permeable AMPA receptors (IC, Petralia and Wenthold, 1992; Martin et al., 1993a; IC, Conti et al., 1994; IC, Kharazia et al., 1996b; IS/IC, Kondo et al., 1997; IC, Petralia et al., 1997). In contrast, most pyramidal neurons have GluR2 but little or no GluR1. Neurons with abundant nitric oxide synthase (NOS) express primarily GluR1 and GluR4 (IS, IC, Catania et al., 1995). Major kainate receptor subunits in the cortex are GluR7 and KA2, although low levels of the other types are found (IS, Wisden and Seeburg, 1993). Immunolabeling for delta 1/2 typically is low (Mayat et al., 1995) (Fig. 2). In the P21 mouse, NR1, NR2A, and NR2B are moderate to high, while NR2C and NR2D may be absent (a faint signal for NR2D, within the range of background, is detected in a small number of small- to medium-sized neurons; IS, Watanabe et al., 1993). In the human cortex, however, NR2C is found in interneurons and NR2D in pyramidal neurons (IS, Scherzer et al., 1998). In the rat, NR2D is found only in a small group of interneurons (IS, Standaert et al., 1996). Neurons with abundant NOS have NR1 variants in which the C1 cassette is absent and the C2' cassette commonly replaces the C2 cassette (IC, Weiss et al., 1998). The latter authors suggest that this particular type of NR1 subunit in these neurons may be responsible for their selective resistance to injury from excess production of NO by NOS elicited by glutamate acting on NMDA receptors. Only low levels of NR3 are found in the adult cortex, although this subunit is abundant during early postnatal development (IS, Ciabarra et al., 1995; IS, Sucher et al., 1995). In the hippocampus, ionotropic glutamate receptors are particularly abundant (Fig. 1). GluR1 and GluR2 have higher levels than GluR3 or GluR4 (IS, Sato et al., 1993a). In contrast to pyramidal cells of the neocortex, both GluR1 and GluR2 are abundant in pyramidal cells of the hippocampus, although like the cortex, many nonpyramidal cells possess GluR1 with little or no GluR2 (IC, Petralia et al., 1997). As in the cortex, neurons with abundant NOS express primarily GluR1 and GluR4 (IS, IC, Catania et al., 1995). Kainate receptors show a variety of patterns: GluR5 is low or absent, GluR6 is low in the CA1/CA3 region and moderate in the dentate gyrus, GluR7 also is moderate in the dentate gyrus but is very low in the CA1/CA3 region, KA1 is high in the dentate gyrus and CA3 region but is nearly absent from the CA1 region, and KA2 is abundant in all areas (IS, Wisden and Seeburg, 1993). Highest levels of delta 1 in the brain are found in the hippocampus (IS, Lomeli et al., 1993) (Fig. 2). Expression of NMDA receptors is similar to that seen in the cortex, with high levels of NR1, NR2A, and NR2B (IS, P21, Watanabe et al., 1993; IC, Petralia et al., 1994a,b). As in the cortex, NR2C 149

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Fig. 2. Sagittal section of rat brain immunolabeled (pre-embedding immunoperoxidase) for delta 1/2. CP = caudate putamen; DC = dorsal cochlear n.; DG = dentate gyms (in hippocampus); FH -- forelimb/hindlimb area of cortex; Fr = frontal cortex; Rt = reticulothalamic n. Note high levels of labeling in the dorsal cochlear nucleus and the cerebellar molecular layer (above DC), moderate labeling in the hippocampus and light to moderate labeling in various other brain structures. Modified from Mayat et al. (1995).

and N R 2 D m a y be a b s e n t in the m o u s e h i p p o c a m p u s (a faint signal for N R 2 D , within the r a n g e of b a c k g r o u n d , is f o u n d in l i m i t e d areas; IS, P21, W a t a n a b e et al., 1993), but are f o u n d in the h u m a n h i p p o c a m p u s (IS, S c h e r z e r et al., 1998). Also, as in the cortex, n e u r o n s with a b u n d a n t N O S t y p i c a l l y have NR1 variants w i t h o u t the C1 c a s s e t t e and c o m m o n l y with a C2' c a s s e t t e (IC, Weiss et al., 1998). N R 1 subunits c o n t a i n i n g the N1 c a s s e t t e are m o r e a b u n d a n t in p y r a m i d a l cells of the C A 2 / C A 3 r e g i o n than in the C A 1 r e g i o n (IS, L a u r i e et al., 1995). In contrast, NR1 subunits c o n t a i n i n g the C1 cassette are a b u n d a n t in the CA1 region but not in the C A 3 r e g i o n (IC, J o h n s o n et al., 1996; Weiss et al., 1998). T h e adult C A 1 r e g i o n c o n t a i n s very low levels of N R 3 a l t h o u g h there is c o n s i d e r a b l e e x p r e s s i o n of N R 3 in early p o s t n a t a l ages (IS, C i a b a r r a et al., 1995; IS, S u c h e r et al., 1995). In the n e o s t r i a t u m ( c a u d a t e p u t a m e n ) , G l u R 1 and G l u R 2 are the m a i n A M P A r e c e p t o r subunits of the m a j o r n e u r o n type m the m e d i u m spiny n e u r o n s (IC, M a r t i n et al., 1993a,b; IS, Sato et al., 1993a; IC 4- s i n g l e - c e l l P C R , C h e n et al., 1998a). G l u R 3 is l o c a l i z e d p r e f e r e n t i a l l y to m e d i u m spiny n e u r o n s c o e x p r e s s i n g s u b s t a n c e P and e n k e p h a l i n (IC 4- s i n g l e - c e l l P C R , Stefani et al., 1998), w h i l e G l u R 4 is e x p r e s s e d p r i m a r i l y in the large c h o l i n e r g i c i n t e r n e u r o n s (IC, M a r t i n et al., 1993b; IC 4- single-cell PCR, Stefani et al., 1998). M o s t k a i n a t e r e c e p t o r s

+

Fig. 1. Coronal sections of rat forebrain immunolabeled (pre-embedding immunoperoxidase) for GluR1 (a), GluR2/3 (b), and GluR4 (c). Note the different patterns: GluR1 is high in some structures and low in others, GluR2/3 is generally high in many structures, and GluR4 is generally low in many structures. Ar -- arcuate hypothalamic n." B1 = basolateral amygdaloid n.; C1 = field CA1 of Ammon's horn; C3 - field CA3 of Ammon's horn; cc = corpus callosum; DG = dentate gyrus; DL = lateral geniculate n., dorsal part; IG - indusium griseum; LA = lateral amygdaloid n." LH -- lateral habenula; LP = lateral posterior thalamic n.; LV -- lateral ventricle; MH - medial habenula; ml = medial lemniscus; PC = posterior cortical amygdaloid n.; P1 = parietal cortex, area 1" Pf = parafascicular thalamic n.; Pi = piriform cortex; rf = rhinal fissure; Rt = reticulothalamic n." St - subthalamic n.; T1 = temporal cortex, area 1" VL = lateral geniculate n., ventral part; ZI = zona incerta; III = third ventricle. From Petralia and Wenthold (1992). 151

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are at low to moderate levels, while GluR6 (IS, Bischoff et al., 1997) and KA2 (IS, Wisden and Seeburg, 1993) may be abundant (IS, Wtillner et al., 1997). Only low levels of delta 1 are found in the adult neostriatum, although higher levels are seen at birth (IS, Lomeli et al., 1993). P21 mice express the NMDA receptor subunits NR1, NR2A and NR2B (IS, Watanabe et al., 1993); in addition to these, rats have low levels of NR2C in some neurons and maybe in glia (IS, Standaert et al., 1999). Cholinergic interneurons show a unique pattern, i.e., NR1 + NR2B + NR2D (IS, Standaert et al., 1996, 1999). As in the cortex and hippocampus, neurons with abundant NOS have NR1 variants in which the C1 cassette is absent and the C2' cassette replaces the C2 cassette (IC, Weiss et al., 1998). Interestingly, NR1 variants containing the N1 cassette are absent from the neostriatum and associated structures (excepting the subthalamic nucleus; IS, Standaert et al., 1994). NR3 is absent from the neostriatum (IS, Sucher et al., 1995). In the amygdala, GluR1-3 are high and GluR4 is low in most nuclei (IS, Sato et al., 1993a) (Fig. 1); GluR4 labeling may be partly or entirely glial (IC, Martin et al., 1993a). Preferential labeling for GluR1 is found in populations of nonpyramidal neurons (IC, Farb and LeDoux, 1997; IC, McDonald, 1996). The main kainate receptor subunit in the amygdala is GluR5 (Hollmann and Heinemann, 1994; Li and Rogawski, 1998). The amygdala of the P21 mouse contains the NR1, NR2A, and NR2B NMDA receptor subunits (IS, Watanabe et al., 1993). Significant levels of NR3 are found in the amygdala (IS, Ciabarra et al., 1995). AMPA receptors vary greatly among thalamic nuclei. Many nuclei express mainly GluR1, GluR2, and GluR4, while others (ventral and lateral groups) express mainly GluR3 and GluR4 (IS, Sato et al., 1993a). GluR4 also is the most common, or the only, AMPA receptor subunit in the reticular nucleus (IC, Petralia and Wenthold, 1992; IC, Martin et al., 1993a; IS, Sato et al., 1993a; IS, IC, Jones et al., 1998) (Fig. 1). Among the kainate receptors, GluR7 is expressed prominently in the reticular nucleus, while GluR5 is prominent in the anteroventral nucleus and is expressed in a unique pattern in a number of small subnuclei near the midline; other kainate receptor subunits are found only in low amounts (IS, Wisden and Seeburg, 1993). Highest levels of delta 1 in the thalamus are found in the anteroventral nucleus (IS, Lomeli et al., 1993). NMDA receptors show varying distributions in the different nuclei of the thalamus. In the monkey but not in the mouse, NR2D is particularly high in the anterodorsal nucleus (IS, P21, Watanabe et al., 1993; Jones et al., 1998). Significant levels of NR3 are found in the thalamus (IS, Ciabarra et al., 1995). Immunolabeling for AMPA receptors is found in the pineal gland of a primate, the cynomolgus macaque, although labeling appears to be absent from the pinealocytes (Mick, 1995). In the rat, substantial immunolabeling of the pineal gland is found with antibodies to KA2 and NR2A/B; somewhat lower levels are found with antibodies to GluR6/7 and NR1 (Petralia et al., 1994a,b,c). mRNA for KA2 (Wisden and Seeburg, 1993) and NR2C (T611e et al., 1993) are expressed abundantly in the pineal gland. Immunolabeling for delta 1/2 is high in the pineal gland (Mayat et al., 1995). Glutamate receptors in the hypothalamus have been reviewed in detail in our previous work (Petralia and Wenthold, 1996). GluR1 and GluR2 are the major AMPA receptor subunits (IS, Sato et al., 1993a; IS, Van den Pol et al., 1994; IC, Ginsberg et al., 1995). The major kainate receptor subunits are GluR5, GluR7 and KA2 (IS, Wisden and Seeburg, 1993). Changes in the expression of KA2 in gonadotropin-releasing hormone (GnRH) neurons occur during sexual maturation in the female rat, indicating that kainate receptors play an important role in regulation of postnatal sexual development (IS, Eyigor and Jennes, 1997). Only low levels of immunolabeling for delta 1/2 are found in the hypothalamus, with densest labeling in the supraoptic nucleus (Mayat et al., 1995). Hypothalamic nuclei show varying combinations of 152

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NMDA receptor subunits. Notable is the high level of NR2C in the suprachiasmatic nucleus (IS, P21, Watanabe et al., 1993). GnRH neurons in the female rat show high levels of NR2A but apparently no NR1; unlike KA2 receptors (see above), NR2A levels do not change during sexual maturation (IS, Eyigor and Jennes, 1997). Some NR3 is found in the hypothalamus (IS, Ciabarra et al., 1995). AMPA, kainate, delta and NMDA receptors are found in varying amounts in the pituitary gland, as discussed in detail in Petralia and Wenthold (1996). 2.2. MID/HINDBRAIN Moderate to high levels of GluR1, GluR2 and GluR4, and low levels of GluR3, are found in the substantia nigra (IS, Sato et al., 1993a). Dopaminergic neurons (i.e., tyrosine hydroxylase-positive) of the pars compacta contain GluR1 and GluR2/3 in the rat (IC, Martin et al., 1993a), and also GluR4 in the monkey (IC, Paquet et al., 1997). In the mouse, the pars compacta contains all kainate receptor subunits except KA1; of these, GluR5 and GluR7 are abundant (IS, B ischoff et al., 1997). Only GluR5 and GluR6 are found in the pars reticulata of the mouse. In contrast, only KA2 and GluR7 in the pars compacta, and KA2 and GluR6 in the pars reticulata, are reported for the rat (IS, Wfillner et al., 1997). There are moderate levels of NR1 and low levels of NR2A and NR2B in the substantia nigra (IS, P21, Watanabe et al., 1994a). Dopaminergic neurons of the pars compacta have NR1 but appear to lack NR2A/B (Paquet et al., 1997) and functional NMDA receptors (Wu and Partridge, 1998). In the cerebellum, Purkinje cells express mainly GluR2 and GluR3, while granule cells express GluR2 and GluR4 (IC, Petralia and Wenthold, 1992; IC, Martin et al., 1993a; IS, Sato et al., 1993a; IC, Zhao et al., 1997). Small neurons of the molecular layer (stellate -+- basket cells) contain GluR2, GluR3 and maybe some GluR4 (IC, Martin et al., 1993a; IS, Sato et al., 1993a; IC, Petralia et al., 1997). Golgi cells express mRNA for GluR3 (Sato et al., 1993a) and immunolabeling for GluR2/3 (Martin et al., 1993a; Petralia et al., 1997). Unipolar brush cells immunolabel moderately for both GluR2 and GluR2/3 (Petralia et al., 1997) and it has been suggested that these neurons contain only homomeric GluR2 (Jaarsma et al., 1995). Of the kainate receptors, GluR5 and KA1 are expressed in Purkinje cells, while GluR6 and KA2 are expressed in granule cells (IS, Wisden and Seeburg, 1993). Stellate/basket cells may contain only GluR7 (IS, Wisden and Seeburg, 1993; Petralia et al., 1994c). Delta 2 is expressed at very high levels in Purkinje cells (IS, Araki et al., 1993; IS, Lomeli et al., 1993; IC, Mayat et al., 1995) (Fig. 2). NR1 is prevalent in both Purkinje cells and granule cells (IS, Akazawa et al., 1994; IS, Watanabe et al., 1994b); it also is present in small neurons of the molecular layer and in Golgi cells (IS, Akazawa et al., 1994); IC, Petralia et al., 1994a) (Fig. 3). In the rat, Purkinje cells also possess small amounts of NR2A, but none of the other NR2 subunits. This pattern is consistent with the apparent absence of functional NMDA receptors in adult Purkinje cells, since most Purkinje cell NMDA receptor complexes would lack the NR2 subunits, which are necessary for normal function (discussed in Petralia et al., 1994a). In both rats and mice, granule cells have NR2A and NR2C (IS, Akazawa et al., 1994; IS, Watanabe et al., 1994b). NR2D is found in Golgi cells and possibly in stellate cells (IS, Akazawa et al., 1994). Strangely, immunolabeling for NR2A/B is found in the pinceau (a highly modified basket cell axon surrounding the proximal portion of the Purkinje cell axon; Petralia et al., 1994b). Since NR2A or NR2B do not appear to be expressed in basket cells, the NR2A/B antibody likely recognizes a type of potassium channel that has an antigenic site similar to that of NR2A or NR2B, and is found in high concentration in the pinceau (discussed in Petralia and Wenthold, 1999). The long variant of NR3 predominates over the short variant 153

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Fig. 3. Sagittal section of the cerebellar cortex immunolabeled (pre-embedding immunoperoxidase) for NR1. Labeling is found in most or all neurons, including Purkinje cells (Pj), granule cells (Gr), Golgi cells (Go), and small cells of the molecular layer (arrowheads). Arrows, Purkinje cell dendrites. Modified from Petralia et al.

(1994a).

in the adult cerebellum; levels of both variants are considerably higher at birth (Sun et al., 1998). In the vestibular nuclei, GluR2 and GluR3 are the predominant AMPA receptor subunits in the rat (Sato et al., 1993a) and chinchilla (Popper et al., 1997). There is little information on the kainate receptors of the vestibular nuclei, although low to moderate levels of labeling are found with antibodies to GluR6/7 and KA2 (Petralia et al., 1994c). All vestibular nuclei of the mouse express some NR1 and NR2A, while none expresses NR2B and some express low levels of NR2D; NR2C is found only in the medial nucleus (IS, P21, Watanabe et al., 1994a; for rat, see IS, De Waele et al., 1994 and IC, Petralia et al., 1994a,b). Expression in the guinea pig is similar but includes NR2B in the medial and lateral nuclei and NR2C in the lateral 154

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nucleus (IS, Sans et al., 1997). NR3 appears to be absent from most parts of the hindbrain (IS, Sucher et al., 1995). In the cochlear nuclei, GluR2-4 are widespread, while GluR1 is found mainly in cartwheel/stellate type cells of the dorsal cochlear nucleus and in a few neurons scattered in the acoustic striae (IS, Hunter et al., 1993; IC, Petralia et al., 1996; IC, Wang et al., 1998). Little is known about kainate receptors in the cochlear nuclei. Low to moderate levels of labeling are found with antibodies to GluR6/7 and KA2 (IC, 1,etralia et al., 1994c; IC, Petralia et al., 1996). High levels of immunolabeling for delta 1/2 are found in the dorsal cochlear nucleus (IC, Mayat et al., 1995; IC, 1,etralia et al., 1996) (Fig. 2). With KA2 antibody, labeling is higher in the large neurons of the cochlear root nucleus, than in other neurons of the cochlear nuclei. NR1 and NR2A are found in most areas of the P21 mouse cochlear nuclei, while distributions of NR2B and NR2C are more restricted, and NR2D seems to be absent (IS, P21, Watanabe et al., 1994a). However, in the rat cochlear nuclei, low levels of NR2D are found throughout. Also, overall highest levels of NR2A-C are found in the small cell cap overlying the anteroventral cochlear nucleus in the rat (IS, Sato et al., 1998). In neurons of the nucleus of the tractus solitarius, GluR1 and GluR2 are the major AMPA receptor subunits (IC, Petralia and Wenthold, 1992; IS, Sato et al., 1993a; IC, Ambalavanar et al., 1998). Low to moderate levels of labeling are found with antibodies to GluR6/7 and KA2 (IC, Petralia et al., 1994c). NR1 and low levels of NR2A and NR2B are found in this nucleus (IC, Petralia et al., 1994a; IS, 1,21, Watanabe et al., 1994a; IC, Ambalavanar et al., 1998). 2.3. SPINAL CORD AND PERIPHERAL In the spinal cord, highest labeling for GluR1 and GluR2 are in the upper dorsal horn, while highest labeling for GluR3 and GluR4 are in the lower dorsal horn and in the motor neurons of the ventral horn (IS/lumbar segments, T611e et al., 1993; IS, IC/mainly cervical segments, other segments mentioned, Furuyama et al., 1993; IC/all segments, Petralia et al., 1997). Overall, labeling for GluR1 is low in the spinal cord (IS, T611e et al., 1993) but is high in a small number of elongate neurons found in laminae X and scattered in other laminae (IC/cervical segments, Martin et al., 1993a; IC/all segments, Tachibana et al., 1994). Populations of GluRl-selective cells in the upper spinal cord (IC/cervical/lumbar segments? Popratiloff et al., 1996; see also Petralia et al., 1997) have calcium-permeable AM1,A receptors (i.e., lacking GluR2; IC/lumbar segments, Engelman et al., 1999). Highest expressed kainate receptors in the spinal cord are KA2 in the upper dorsal horn and KA1 in the motor neurons of the ventral horn; GluR5-7 are low or absent (IS, T611e et al., 1993). mRNAs for delta 1 and delta 2 are only weakly detectable in the spinal cord, although delta 1 is slightly higher in the motor neurons of the ventral horn (IS, T611e et al., 1993). NR1 and NR2A are widespread in the spinal cord of the mouse; in addition, a low level of NR2B is found in the upper dorsal horn, while no NR2C or NR2D are seen (IS/cervical segments, Watanabe et al., 1994c). Some NR3 is found in the spinal cord, with highest levels in laminae 2-3 (IS/cervical/lumbar segments, Ciabarra et al., 1995). Dorsal root ganglion neurons express mainly labeling for GluR2/3, while the associated satellite cells (a type of glia) label densely for GluR4 (IC, Sato et al., 1993b; IC, Tachibana et al., 1994) (Fig. 4). GluR5 is expressed strongly in the small ganglion neurons (IS, Sato et al., 1993b). Immunolabeling for delta 1/2 is moderate in ganglion cells (IC, Mayat et al., 1995). NR1 is expressed in all neurons (IS, Sato et al., 1993b). In the 1'21 mouse, NR1 is expressed in both the dorsal root and trigeminal ganglia whereas the NR2 subunits are absent (IS, Watanabe et al., 1994d). 155

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Fig. 4. Cervical dorsal root ganglia immunolabeled (pre-embedding immunoperoxidase) for GluR2/3 (a) and GluR4 (b). Arrows indicate satellite cells showing little or no immunolabeling with antibody to GluR2/3 and stained densely with antibody to GluR4. From Tachibana et al. (1994), reproduced with permission from Wiley-Liss, Inc.

In the superior cervical (sympathetic) and pterygopalatine (parasympathetic) ganglia, labeling with GluR2/3 antibody is prevalent; labeling with GluR1 antibody is about half as common and labeling with GluR4 antibody is limited to a group of small, specialized neurons (small, intensely fluorescent, SIF, cells) (IC, Kiyama et al., 1993). Immunolabeling for GluR1, GluR2/3 and GluR4 are found in the submandibular ganglion and in associated structures of the salivary glands, while NR1 immunolabeling is absent (Shida et al., 1995). Cochlear and vestibular ganglia express substantial GluR2-5 and NR1, and low to moderate amounts of GluR6, NR2A-D and KA1-2, while GluR1 and GluR7 are absent (IS, Safieddine and Eybalin, 1992; IC, Kuriyama et al., 1994; IS, Niedzielski and Wenthold, 1995). Delta 1 also is very prevalent in cochlear and vestibular ganglia and in satellite cells of the cochlear ganglion (IS, IC, Safieddine and Wenthold, 1997). Details about glutamate receptors in the organ of Corti are given in Chapter IX, by Usami et al. Immunolabeling for GluR1, GluR5/6/7 and NR1 is found in unmyelinated axons in glabrous skin of the rat hindpaw (Carlton et al., 1995). They are believed to act as autoreceptors for secreted glutamate that may regulate the response to pain. A number of organs have glutamate receptors. In the adrenal gland, (1) GluR1 and GluR3 predominate in different parts of the cortex, (2) GluR2 is in medullary cells, (3) GluR4 156

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is present only in very low levels, and (4) all four AMPA receptor subunits are found in medullary ganglion cells; this suggests that different cell populations in the adrenal gland may have different AMPA receptor types, some apparently homomeric (IS, Kristensen, 1993). NR1 is found in the adrenal medulla of the P21 mouse (NR2A-D are absent; IS, Watanabe et al., 1994d). In the pancreas of newborn guinea pigs, central insulin-secreting cells of the islets of Langerhans express GluR1 and GluR4, while the peripheral cells show GluR2/3 labeling, and GluR2/3 plus GluR4 are found in pancreatic ganglion cells (IC, Liu et al., 1997b). Neurons of the enteric nervous system, which innervate the gastrointestinal system, express GluR1-4 differentially in different cell populations, while labeling for NR1 and NR2A/B is common throughout these neurons (IS, Burns and Stephens, 1995; IC, Liu et al., 1997a). Other examples of glutamate receptors in organs are in the respiratory system (Said et al., 1995) and in bone cells. Glutamate receptors are found in peripheral cholinergic nerves that innervate bronchial smooth muscle, and may explain symptoms of 'Chinese restaurant syndrome' due to ingested glutamate (Aas et al., 1989). In bone, NR1 is localized in osteoblasts and osteoclasts (IS, IC, Patton et al., 1998), apparently in association with NR2D and the associated protein, PSD-95 (as determined with PCR; NR2A-C absent); this study suggests that bone cells signal each other via glutamate transmission. Putative glutamate receptors also have been reported in human peripheral monocytes (Malone et al., 1986). Taste bud cells in the mouth are believed to respond to the taste of glutamate through the metabotropic glutamate receptor, mGluR4 (Chaudhari and Roper, 1998). However, there is evidence for ionotropic glutamate receptors, probably NMDA receptors, in taste bud cells (Chaudhari and Roper, 1998). It is not clear whether the latter receptors participate in taste transduction at the apical (tasting) end of the cell or are involved in synaptic transmission at the basolateral (neural) end of the cell. Finally, glutamate receptors are found in developing neuromuscular synapses. In arthropod muscles, glutamate is the major neurotransmitter while acetylcholine is the major one in vertebrates (e.g., Betz et al., 1993). Nevertheless, presynaptic ionotropic glutamate receptors are found in the neuromuscular junctions in lower vertebrates during development and probably regulate neurotransmitter release at this synapse (Fu et al., 1995; Chen et al., 1998b). In addition, postsynaptic NR1 immunolabeling has been described at neuromuscular junctions in mice and rats (Berger et al., 1995; Grozdanovic and Gossrau, 1998). 2.4. RETINA Distribution of glutamate receptors in the retina has been reviewed recently (IS, Brandst~itter et al., 1998). GluR1-4 are expressed in patches of cells within the ganglion cell layer (GCL) and in cells of the inner third (amacrine cell region) of the inner nuclear layer (INL); GluR1 and GluR2 are expressed in almost all cell bodies of the INL while GluR3 and GluR4 show more limited distributions (IS, Mtiller et al., 1992; IS, Hamassaki-Britto et al., 1993). GluR3 is expressed prominently in a subset of large cells (probably horizontal cells) of the outer edge of the INL of the cat but not of the rat (IS, Hamassaki-Britto et al., 1993); these may be type-A horizontal cells which are absent in the rat (Lo et al., 1998). In the cat, type-A horizontal cells label for both GluR2/3 and GluR4, while type-B horizontal cells express only GluR4 (IC, Morigiwa and Vardi, 1999). GluR1 appears to be absent from horizontal cells in the cat (IC, Qin and Pourcho, 1999). GluR5 is expressed in rare cells of the GCL and in the outer two-thirds of the INL, while GluR6 and GluR7 are expressed more commonly in cells of the GCL and are expressed throughout the INL (Hamassaki-Britto et al., 1993). Another study indicates that in the INL, 157

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GluR6 is limited to a subset of amacrine cells, while GluR7 is expressed in most amacrine and bipolar cells (inner and middle INL, respectively), but probably not in horizontal cells (IS, Brandst~itter et al., 1994). KA1 is not expressed in the rat retina, although described in the mouse retina (Zhang et al., 1996); in contrast, KA2 is common in cells throughout the GCL and INL (IS, Brandst~itter et al., 1994). Immunolabeling for delta 1/2 is restricted to the neuropil of the inner plexiform layer (IC, Brandst~itter et al., 1997). NR1 is expressed throughout the GCL and INL, while NR2A and NR2B are expressed throughout the GCL but only in some amacrine cells of the inner INL (IS, Brandst~itter et al., 1994; IS, Hartveit et al., 1994). In contrast, NR2C is expressed throughout the GCL and INL (IS, Brandst~itter et al., 1994); NR2B (s2) also is expressed throughout the INL in P21 mice (Watanabe et al., 1994e). NR2D expression has not been detected with in situ hybridization (IS, Brandst~itter et al., 1994), although Wenzel et al. (1997) report immunolabeling for NR2D in rod bipolar cells; however, these cells appear to lack functional NMDA receptors (see discussion in IS, Brandst~itter et al., 1998). Interestingly, immunolabeling for NR2A and NR2B has been described in rod and cone outer segments, using antibodies shown to be specific in brain tissue (Goebel et al., 1998). An interesting phenomenon is found in the immunolabeling of bipolar cell dyad synapses, which consist of a single presynaptic terminal and two postsynaptic elements. In every case studied, only one of the two postsynaptic elements is labeled, including for GluR1, GluR2/3, GluR6/7, KA2, delta 1/2, NR2A, and the metabotropic receptors, mGluRl~, mGluR5, and mGluR7 (Brandst~itter et al., 1997; Qin and Pourcho, 1999; also equally selective presynaptic localizations for mGluR7 see Brandst~itter et al., 1996). Thus, the bipolar cell terminal presumably is contacted by two postsynaptic elements with different receptor combinations.

3. NEURONAL DISTRIBUTION

3.1. SYNAPTIC DISTRIBUTION 3.1.1. Adult synapses

Glutamate receptors vary in their distributions at synapses (Ottersen and Landsend, 1997; Petralia, 1997; Somogyi et al., 1998; Petralia et al., 1999c,d). AMPA receptors are found most commonly in the postsynaptic membrane, although there is limited evidence for presynaptic AMPA receptors. Kainate receptor distribution at synapses is still not well understood; immunocytochemical data support a mostly postsynaptic localization, although other lines of evidence indicate that kainate receptors may be most common in the presynaptic membrane and in perisynaptic membrane (on the postsynaptic side). Delta receptors are found in the postsynaptic membrane. NMDA receptors are found in the postsynaptic membrane, but there is some evidence for presynaptic NMDA receptors. Metabotropic glutamate receptor distribution at the synapse varies the most. For example, mGluRlc~ and mGluR5 are found mainly in the perisynaptic membrane (on the postsynaptic side), while mGluR7 is best known as a presynaptic receptor. 3.1.1.1. Differential distribution

Both glutamate (Landsend et al., 1997; Rubio and Wenthold, 1997; Zhao et al., 1997, 1998; Toth and McBain, 1998) and GABA (Nusser et al., 1996a,b) receptors have been shown to be 158

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differentially distributed in individual neurons. This is not surprising, considering that most neurons receive numerous different excitatory and inhibitory inputs; presumably the neuron has developed a mechanism to target selectively its multiple receptor types to different populations of synapses, to allow for multiple physiological responses. Differential distribution of glutamate receptors occurs on synapses located either on two separate dendrites (apical versus basal dendrites) or on different regions of the same dendrite; possible differential distributions involving dendrites and cell body excitatory synapses also have been described (Wang et al., 1998). Differential distribution of glutamate receptors in apical versus basal dendrites probably occurs in pyramidal cells of the hippocampus and cerebral cortex, but definitive studies have not yet been done. However, this phenomenon has been studied in detail in the fusiform cell of the dorsal cochlear nucleus (Rubio and Wenthold, 1997). The fusiform cell is a bipolar neuron with apical and basal dendritic trees, receiving two different excitatory synaptic inputs, i.e., parallel fibers from the granule cells on apical dendrites, and the primary input from the auditory nerve on basal dendrites (Fig. 5). The fusiform cell expresses multiple subtypes and subunits of glutamate receptors, including GluR2/3, GluR4, NR2A/B, delta 1/2 and mGluRlc~. By retrograde tracing and postembedding immunogold labeling, fusiform cells were shown to express different glutamate receptors at these two synapse populations (Table 2). Subunits like GluR2/3 and NR2A/B are equally abundant at both synaptic populations, while GluR4 and mGluR1 c~ are present only at the basal dendrite synapses. Delta 1/2 is about 4 times more abundant at apical dendrite synapses. Physiological studies confirm that metabotropic glutamate receptors are present in fusiform cells but they do not modulate responses evoked by parallel fiber stimulation (Molitor and Manis, 1997); this indicates that functional metabotropic glutamate receptors are absent from synapses on apical dendrites. The preferential presence of the GluR4 subunit on auditory nerve synapses and its absence from parallel fiber synapses can be related to the fast rate of desensitization of GluR4 (Mosbacher et al., 1994). AMPA receptors with a high content of GluR4 have fast responses. The rapid firing of the auditory inputs to the basal dendrites of fusiform cells presumably is necessary for accurate sound localization by the auditory nuclei. The other form of differential distribution of glutamate receptors, i.e., in two synapse

TABLE 2. Summary of the postembedding immunoreactivity for glutamate receptor subunits at the auditory nerve and parallel fiber synapses Receptors

GluR2/3 a,b GluR2 b GluR4 b GluR4 (10 nm gold) NR2A/B b mGluRlo~ a'b Deltal/2 b

Auditory nerve synapses (basal dendrites)

Parallel fiber synapses (apical dendrites)

No. PSDs

No. gold particles/ltm of PSD 4- SE

No. PSDs

No. gold particles/Ixm of PSD 4- SE

18 25 17 9 19 35 31

17.7 9.1 19.1 9.2 6.4 8.0 8.3

17 17 17 8 14 25 25

16.5 7.2 0 0 9.8 0 33.9

4- 4.0 4- 1.1 4- 2.2 4- 1.9 4- 1.4 4- 1.3 4- 1.2

4- 3.2 4- 1.2

4- 1.3 4- 3.1

a Monoclonal antibodies. b5 nm gold was used for immunogold-labeling quantification with all the antibodies selective for the glutamate receptor subunits, except for GluR4 which was analyzed using 5 nm and 10 nm. Table modified from table 2 in Rubio and Wenthold (1997).

159

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Fig. 5. Electron micrograph montage of a secondary apical dendrite (ADII) of a fusiform cell of the dorsal cochlear nucleus (DCN) after immunogold labeling (5 nm gold) with a polyclonal antibody to GluR2/3 receptor subunits. Two parallel fiber synapses of the granule cells (1, 2) are observed making synaptic contact on a dendritic spine (1) and the dendritic shaft (2). An electron-dense granule of HRP (arrowhead; used for retrograde tracing) can be seen in the dendrite. (B) Drawing of the same apical dendrite (A) showing the synaptic [postsynaptic membrane of the parallel fiber synapses (1, 2)] and subcellular location of gold particles labeling GluR2/3 subunits. The size of the gold particles has been increased for a better visualization. The lines inside the dendritic profile represent cytoskeleton and membranous structures. The arrow is oriented toward the surface of the DCN and away from the cell body. Scale bar, 2 Ixm. (C) Schematic drawing showing the excitatory synaptic circuit on fusiform cells and the division of the apical and basal dendritic segments. Types and subunits of glutamate receptors expressed at the postsynaptic membrane of the auditory nerve (AN) and parallel fibers of the granule cells (PF) are indicated. From Rubio and Wenthold (1999a).

populations on the same dendrite, has been described for Purkinje cells of the cerebellum and for two kinds of neurons of the CA3 region of the hippocampus. Purkinje cells have two excitatory inputs, i.e., climbing fiber synapses originating from inferior olivary neurons, 160

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0

0

~

Ot Ot

Fig. 6. Summary histogram of development of glutamate receptors at parallel [postnatal day 10 (P10) to adult] and climbing (P2 to adult) fiber synapses on Purkinje cells of the cerebellum. Note especially the peak in immunogold labeling of the delta receptors at P 10-P 14 in climbing fiber synapses (cf), the peaks of the AMPA receptors (GluR2, GluR2/3 antibodies) at P2-P5, and the inverse patterns of peaks for parallel fiber synapses (pf) and climbing fiber synapses in adults for AMPA versus delta receptors. Modified from Zhao et al. (1998).

and parallel fiber synapses originating from granule cells of the cerebellum. Delta 2 receptors are abundant at parallel fiber synapses but are rare or absent from climbing fiber synapses (Landsend et al., 1997; Zhao et al., 1997). In contrast, AMPA receptors (labeled for GluR2/3 or GluR2) are more common in climbing fiber synapses than in parallel fiber synapses (Zhao et al., 1998) (Fig. 6). Delta 2 is believed to play a specific role in synaptic plasticity of adult parallel fiber synapses, since long-term depression of parallel fiber synapses is impaired in knockout mice lacking delta 2 (Kashiwabuchi et al., 1995). In the apical dendrites of pyramidal cells of the CA3 region of the hippocampus, postsynaptic immunolabeling for NR1 subunits is more common at small spine synapses than at mossy terminal synapses (Petralia et al., 1994a; Siegel et al., 1994). This was shown also with immunogold labeling using a mixture of NR1 and NR2A/B antibodies (Takumi et al., 1999). In contrast, immunolabeling for NR2A/B is abundant in at least some mossy terminal synapses (Petralia et al., 1994b). Studies using separate antibodies for NR2A and NR2B suggest that, in CA3 pyramidal cell apical dendrites, NR2A is present in small spine and mossy terminal synapses, while NR2B is present in small spine synapses and absent from mossy terminal synapses (Fritschy et al., 1998; also Watanabe et al., 1998). Thus, NMDA receptor composition may differ between small spine and mossy terminal synapses in CA3 pyramidal cell apical dendrites. In support of this, most apical dendrite synapses exhibit NMDA-receptor-dependent long-term-potentiation (LTP), while mossy terminals have NMDA-receptor-independent LTP (Zalutsky and Nicoll, 1990; Derrick et al., 1991), even though they do have some functional NMDA receptors (Spruston et al., 1995). Differential distribution of NMDA receptors with different subunit compositions also is supported by developmental studies (see Section 3.1.2). In the CA3 region, differential distribution also occurs in interneurons, which have calcium-permeable AMPA receptors (lacking GluR2?) at mossy fiber terminals and calcium-impermeable AMPA receptors at commissural/associational axon terminals (Toth and McBain, 1998). Also, different populations of AMPA receptors, containing GluR1 plus GluR2, GluR2 plus GluR3, or GluR1 161

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only, are found in the CA1/CA2 region, although their synaptic distribution is not known (Wenthold et al., 1996). In addition, some synapses in the hippocampus and other regions may have NMDA receptors but lack AMPA receptors, while most synapses on the same neurons have both NMDA and AMPA receptors. The former synapses are called 'silent synapses' and can acquire AMPA receptors following adequate activation (e.g., review by Malenka and Nicoll, 1997; Nusser et al., 1998; Petralia et al., 1999a; Shi et al., 1999) (see Section 3.1.2). 3.1.1.2. Tangential distribution

Often the distribution of glutamate receptors along the length of the postsynaptic membrane (i.e., the tangential distribution) appears to vary. This has been observed mainly for AMPA receptors which may be more common in the outer portions of the postsynaptic membrane, as noted in the cerebral cortex (Kharazia et al., 1996b; Kharazia and Weinberg, 1997), neostriatum (Bernard et al., 1997), cerebellum (Petralia et al., 1998), and cochlea (Matsubara et al., 1996). Some evidence also exists for restricted tangential distribution of NMDA receptors, (Kharazia et al., 1996a; Kharazia and Weinberg, 1997; Somogyi et al., 1998, Racca, et al., 2000). Assuming that these differences in tangential distribution of glutamate receptors are significant, they may reflect either an adaptation to release of neurotransmitter or a regulation of receptor numbers. In the former case, differences in tangential distribution of glutamate receptors may be related to the position of one or multiple release sites (Harris and Sultan, 1995), as discussed in Matsubara et al. (1996) and Xie et al. (1997). In the latter case, differences in tangential distribution, in particular the greater abundance of receptors along the outer portion of the postsynaptic membrane, may reflect movement of receptors to and from the synapse (see below). Finally, there is a more definitive difference in tangential distribution between postsynaptic ionotropic and some metabotropic glutamate receptors. mGluRl~ and mGluR5 are found mainly in the perisynaptic region of synapses (Baude et al., 1993; Luj~in et al., 1996, 1997; Petralia et al., 1998) while ionotropic glutamate receptors are uncommon in the perisynaptic region (Nusser et al., 1994; Petralia et al., 1998). Preferential distribution of metabotropic glutamate receptors to the perisynaptic region may keep these receptors at a certain distance from the neurotransmitter release sites in the terminal, so that metabotropic glutamate receptors will respond only when a large quantity of glutamate is released. 3.1.1.3. Synaptic zones

We suggest that the synaptic spine can be divided into four major zones containing different combinations of proteins. This idea is based on analyses of numerous glutamate receptors and associated proteins, using postembedding immunogold of fixed or live tissue from the hippocampus and cerebellum (Petralia et al., 1999b). It was found that each associated protein typically has a preferential localization in one zone of the spine synapse, although labeling for the protein may be present to a lesser extent in the other zones (Fig. 7). Our fixed tissue technique has been described in several papers (Rubio and Wenthold, 1997, 1999a; Petralia et al., 1997, 1998, 1999a,b; Wang et al., 1998; Zhao et al., 1998; for a detailed description, see Petralia and Wenthold, 1999); it was based originally on the methods of Matsubara et al. (1996) and Landsend et al. (1997). In the live tissue technique (unpublished data in Petralia et al., 1999b; modified from a similar method described in Petralia and Wenthold, 1998), the live tissue is removed from the brain quickly, slam-frozen, and stored 162

Regional and synaptic expression of ionotropic glutamate receptors

I~ ]t~

mGluRs

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0...00

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Fig. 7. Schematic diagram illustrating how glutamate receptors may be incorporated into, arranged in, and removed from the postsynaptic spine. Glutamate receptors are incorporated into the membrane of vesicles or into the continuous reticulum (gray shading) of the dendrite. Movement of the receptors is controlled by motor molecules such as kinesins on microtubules in the dendrite shaft, and myosins on actin filaments in the spine. Vesicles carrying glutamate receptors may exocytose along the dendrite shaft, allowing the receptors to diffuse up into the spine and ultimately into the postsynaptic membrane, or the vesicles may exocytose along the side of the spine head and then diffuse into the postsynaptic membrane. Alternatively, receptors may reach the postsynaptic membrane by traveling along the continuous reticulum, which may form bridges contacting the postsynaptic membrane and density. Retention of ionotropic glutamate receptors at the postsynaptic membrane may involve various anchoring proteins, particularly of the PSD-95 family, as well as some anchoring proteins specific for AMPA receptors. Various cytoskeletal proteins may be involved in maintaining these protein complexes at the postsynaptic membrane, for example, associations involving ~-actinin and actin. In addition, many proteins including those of the PSD-95 family may be involved in secondary pathways that transduce the postsynaptic signal. Some glutamate receptors, mainly kainate and metabotropic glutamate receptors (mGluRs), are found outside of the postsynaptic membrane in the perisynaptic or presynaptic membranes. See text for details. Based on similar diagrams in Petralia et al. (1999c,d).

in liquid nitrogen. Then, it is p l a c e d directly in the freeze-substitution apparatus so that all fixation, washing and cryoprotection steps are eliminated. Thus, the live t i s s u e - p o s t e m b e d d i n g technique avoids several steps that m a y lead to artefacts in distribution. This m e t h o d also results in increased i m m u n o l a b e l i n g in m a n y cases, and should provide a m o r e realistic view of the postsynaptic density (PSD). The first of the four zones includes the postsynaptic m e m b r a n e and a p p r o x i m a t e l y the upper half of the PSD. In this zone, preferential labeling is seen for all ionotropic g l u t a m a t e receptors and for the associated proteins k n o w n as m e m b r a n e - a s s o c i a t e d guanylate kinases ( M A G U K s ) including PSD-95, SAP-102, PSD-93, and SAP-97 (Valtschanoff et al., 1999) (Figs. 7 and 8). The second zone includes a p p r o x i m a t e l y the b o t t o m half (for the purpose 163

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Fig. 8. Immunogold labeling of AMPA receptors in the CA1 stratum radiatum of the hippocampus. Postembedding immunogold labeling was done using antibodies to GluR1 C-terminus (a, d, g) and GluR2/3 (b, c, e, f, h, i) with 10 nm gold, at postnatal day 2 (P2; a-c), postnatal day 10 (P10; d-f), and 5 weeks (g-i). p = presynaptic terminal. Line scale is 0.2 g m . Micrographs were chosen to illustrate the major trend, that is, a large increase in labeling for AMPA-Rs at 5 weeks compared to P2/P10 (a-f versus g-i). From Petralia et al. (1999a).

of orientation only) of the PSD and a thin region just subjacent to the PSD. The structural boundaries of this zone are not completely clear although our studies with live postembedded tissue suggest that one boundary may center on the postsynaptic lattice that is believed to form the bottom edge of the PSD (Matus and Taft-Jones, 1978; Bloch et al., 1997; Ziff, 1997). Proteins common in this zone include homer lb,c and homer 3 (proteins associated with metabotropic glutamate receptors), CRIPT and GKAP (proteins associated with PSD-95; Naisbitt et al., 1997; Niethammer et al., 1998), CAM-kinase II, and SHANK (a protein associated with both GKAP and homer; Naisbitt et al., 1999; Tu et al., 1999). SynGAP (a protein associated with PSD-95 and SAP-102; Kim et al., 1998) seems to be equally common in zones one and two; SAP-102 also is common in both zones. The third zone includes the tubulovesicular network in the spine head; in places, this can make direct contact with the bottom or the side of the PSD (see below). Proteins common in this region include homer 2, GRIP (a protein associated with AMPA receptors), IP3 receptors, dynein light chain (Naisbitt et al., 2000), and myosin V. Interestingly, homer 3 and CRIPT seem to be 164

Regional and synaptic expression of ionotropic glutamate receptors

Ch. V

common in the third zone using the live tissue-postembedding technique, but not with the fixed tissue-postembedding technique; this may be due to a rearrangement of the tissue during fixation or may indicate a change in availability of the antigenic sites. Actin is common in this zone and it accompanies the tubulovesicular structures; actin is found only occasionally in the first two zones. Finally, the fourth zone includes the perisynaptic (near the PSD) and extrasynaptic or nonsynaptic (the remainder of the spine head) cell membrane. Of the proteins studied, the only ones preferentially localized to this zone are the metabotropic glutamate receptors, mGluRl~ and mGluR5, as noted above. The preferential positions of these various proteins in the four zones can be due to any combination of the following three reasons. (1) The protein has a cytoskeletal role, i.e., it is fixed in place and supporting the localization of other proteins. The best examples of this are the MAGUKs such as PSD-95, which may anchor NMDA receptors, and actin, which, in addition to its cytoskeletal functions, may form the major track for the movement (at least within spines) of other proteins along the reticulum or in vesicles (Ziff, 1997; Tabb et al., 1998). (2) The protein is moving. For example, SAP-102 in the second zone (deep PSD) may be in the process of moving to the first zone where it would have a cytoskeletal function. It could be moving alone or could be involved in the transport of glutamate receptors or other proteins. Another example is myosin V that could move other proteins along the actin pathways (Mermall et al., 1998; Tabb et al., 1998). Proteins may be positioned for short distance movements as part of a transduction mechanism. For example, the dual localization of synGAP in zones 1 (upper PSD) and 2 (deeper PSD) may reflect the position of synGAP molecules that are attached to PSD-95 in the upper zone and are modulating Ras GTPase activity (Kim and Huganir, 1999) in the deeper zone. (3) The protein is being stored for future use. For example, some proteins in zones 2 or 3 may be stored for future use in zone 1. It has been suggested that AMPA receptors may be stored in reticular structures of the spine (zone 3) and in the nonsynaptic (or extrasynaptic) cell membrane of hippocampal spine synapses (Nusser et al., 1998), but it is not clear whether this represents a store separate from that in the dendrite shafts (Rubio and Wenthold, 1999a; see below).

3.1.2. Developing synapses Regional differences in ionotropic glutamate receptor development will not be covered here in detail; see reviews by Bahn and Wisden (1997) and Watanabe (1997). Many changes in glutamate receptor distribution occur in the first three to four weeks of postnatal development in rodents. Some ionotropic glutamate receptors are more common overall in early postnatal ages and decrease in adults. Examples include delta 1, NR2B, NR2D, and NR3A. In the developing cerebellum, NR2B is replaced by NR2A and NR2C. NR2B appears to perform specific functions in early postnatal development, so that mutant mice lacking NR2B (Kutsuwada et al., 1996) or expressing NR2B without an intracellular C-terminal domain (Sprengel et al., 1998) die around birth. The AMPA receptor subunit, GluR1, increases during development in the neocortex and hippocampus and decreases during development in the striatum and in the granule and Purkinje cells of the cerebellum (Martin et al., 1998). In some auditory brainstem nuclei, GluR1 and GluR2 are highest in early postnatal development, while GluR4 develops later, when GluR1 and GluR2 are decreasing (Caicedo and Eybalin, 1999). In the adult cerebellum, as noted above, parallel fiber synapses have abundant delta receptors, while delta receptors are rare or absent from climbing fiber synapses. AMPA receptors are found at both excitatory synapse populations but are more abundant at climbing fiber synapses. In the first postnatal week, presumptive climbing fiber synapses have high 165

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levels of AMPA receptors while delta receptors are low (Takayama et al., 1996; Zhao et al., 1997, 1998) (Fig. 6). Thus, AMPA receptors are established early at synapses in the cerebellum. By the second postnatal week, parallel fiber synapses are forming; at this time, delta receptors are abundant in both parallel and climbing fiber synapses (while AMPA receptors remain common) (Fig. 9). In addition, the adult dendrites are formed and climbing fiber innervation of Purkinje cells is reduced from multiple climbing fibers to a single climbing fiber with multiple synapses. Delta receptors are concentrated at parallel fiber synapses and at climbing fiber synapses that innervate the new Purkinje cell dendrite; they remain low at somal climbing fiber synapses that are destined to be lost. Delta receptors also are absent from GABAergic synapses (Fig. 9), indicating that, in the second postnatal week, Purkinje cells already have developed differential targeting mechanisms (see below) for delta receptors. Since delta receptors are abundant at both parallel and climbing fiber synapses in the second postnatal week but are abundant only in parallel fiber synapses in adults, they may have one function in the formation of adult parallel and climbing fiber synapses and a different function, specific to parallel fiber synapses, in adults. Indeed, this has been suggested by experimental studies (Kashiwabuchi et al., 1995; Kurihara et al., 1997). In contrast to the situation in the cerebellum, AMPA receptors are uncommon in synapses of the CA1 stratum radiatum region of the hippocampus in early postnatal times (postnatal days 2 and 10; Petralia et al., 1999a) (Fig. 8). At these times, NMDA receptors are found at moderate levels and show only a modest increase in adults. Thus, many synapses at postnatal days 2 and 10 contain NMDA receptors but lack AMPA receptors. Presumably, such synapses correspond to the 'silent synapses' observed in physiological studies (Durand et al., 1996; Wu et al., 1996; Malenka and Nicoll, 1997). Such synapses probably are not found on Purkinje cells, where AMPA receptors are abundant at early synapses. Thus, at least two major patterns of early synapse development are indicated by these studies the NMDAR +/AMPAR'silent synapse, type of the hippocampus and the AMPAR + type seen in Purkinje cells. The distribution and function of NMDA receptors in Purkinje cell synapse development is not well understood. At birth, Purkinje cells have NR1 along with NR2B (mice; Watanabe et al., 1994b) or NR2D (rats: Akazawa et al., 1994); adults have high levels of NR1 and either no NR2 subunits (mice) or NR2A only (rats). Physiological studies have yielded mixed results (discussed in Petralia et al., 1994a). It is likely that Purkinje cells have functional NMDA receptors at least at early postnatal times and that they play roles in synapse maturation (Rabacchi et al., 1992; Vallano et al., 1996). In vitro studies confirm that there are at least two major developmental sequences of glutamate receptor acquisition in neurons. In cultured rat spinal cord, AMPA receptors cluster at immature synapses, probably independent of NMDA receptors (Mammen et al., 1997; O'Brien et al., 1997). In hippocampal cultures, two sequences are seen. In spines, AMPA receptors form the first clusters and colocalize with NMDA receptors later (Rao et al., 1998). In contrast, on dendrite shafts, NMDA receptors form early synaptic and nonsynaptic clusters that are believed to lack AMPA receptors. If true, then these would be 'silent synapses' as described in vivo in the hippocampus. The presence of AMPA-receptor-first synapses in the cerebellum in vivo and in the spinal cord and hippocampus in vitro suggests that this is the major developmental pattern in the brain. The presence of NMDA-receptor-first synapses in the hippocampus in vivo and in vitro suggests that a second developmental sequence may have evolved in higher brain centers. In vitro studies also confirm that some specific targeting mechanisms are present in young neurons, as noted above for the in vivo studies of the cerebellar Purkinje cells. Thus, in hippocampal cultures, AMPA and GABA receptors cluster independently at glutamatergic and GABAergic synapses, respectively (Craig et al., 166

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Fig. 9. Colocalization of GABA (30 nm gold) neurotransmitter and delta 1/2 receptors (10 nm gold) at postnatal day 10. Note the absence of GABA labeling in parallel fiber (pf; a, b) and climbing fiber (cf; b) terminals and the abundant labeling for GABA in Purkinje cell dendrites (P; b) and somata (P; c, d) and in pleomorphic vesicle-containing synaptic terminals (i). Delta receptor labeling is abundant in the postsynaptic density/membrane of parallel and climbing fiber synapses (arrowheads) but is absent from GABAergic synapses (arrows). Scale bar, 0.5 I~m. From Zhao et al. (1998).

1994; Rao et al., 1998). Also and as discussed below, formation of postsynaptic structures at glutamatergic synapses may require an initial cluster of glutamate receptor-associated proteins that can anchor the glutamate receptors to the postsynaptic membrane. 167

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In addition to developmental changes in combinations of receptor types at synapses, changes also occur in subunit compositions of receptor complexes. An interesting model has been suggested for the development of NMDA receptors, based on in vitro hippocampal neuron cultures (Tovar and Westbrook, 1999). A young neuron may have NR1/NR2B receptors in extrasynaptic sites and at immature synapses. Then as the synapses mature, NR1/NR2A/NR2B receptors would be targeted to these synapses. 3.2. CYTOPLASMIC DISTRIBUTION It now is well established that many ionotropic glutamate receptors are abundant in cytoplasm of neuron cell bodies and dendrites. In cultures from hippocampus, cerebellar granule cells, and spinal cord, only about half of the NR1 protein is expressed on the surface, while surface expression of NR2 is more common (Hall and Soderling, 1997a; Mammen et al., 1997; Huh and Wenthold, 1999). Similarly, only about half of total AMPA receptors are found on the surface of cultured hippocampal neurons (Hall and Soderling, 1997b). In living, cultured hippocampal neurons, AMPA receptors tagged with green fluorescent protein are evident throughout the dendrite (Doherty et al., 1997). Cytoplasmic glutamate receptors have been demonstrated with immunoperoxidase and immunofluorescence techniques both in vivo (e.g., Petralia and Wenthold, 1992; Martin et al., 1993a; Petralia et al., 1994a,b, 1997; Petralia, 1997; Zhao et al., 1997) and in vitro (Mammen et al., 1997). At the ultrastructural level, immunoperoxidase labeling can be seen in the cytoplasm in distinct patterns with patches of staining associated with endoplasmic reticulum (ER), Golgi apparatus, and mitochondria (Eshhar et al., 1993; Kharazia et al., 1996a). Associations with mitochondria usually are localized to one pole of the mitochondrion, where the staining is associated with a reticular structure (reviewed in Petralia, 1997; see also Rizzuto et al., 1998). We used immunogold labeling to localize the intracellular pool of glutamate receptors in fusiform cells of the dorsal cochlear nucleus, Purkinje cells of the cerebellum and pyramidal cells of the hippocampus (Rubio and Wenthold, 1999a). We found that groups of gold particles labeling AMPA and metabotropic glutamate receptors are associated with tubulovesicular membranes of the endoplasmic reticulum and cytoskeleton (Figs. 5 and 10). This association starts at the level of the trans-Golgi network in the cell body and continues further throughout dendrites and dendritic spines. It has been shown previously that membranes of the ER extend from the cell body to the most distal dendrites (Walton et al., 1991; Terasaki et al., 1994), including dendritic spines (Spacek and Harris, 1997), and that such membranes express ER and Golgi proteins (Gardiol et al., 1998; Jareb and Banker, 1998; Rubio and Wenthold, 1999b). The presence of these proteins in dendrites has been related to the local synthesis of some proteins (e.g., the ~1 subunit of the glycine receptor), but it also indicates that proteins synthesized in the cell body can undergo posttranslational processing, such as glycosylation and assembly, in the dendrite (Rubio and Wenthold, 1999b). We find that the intracellular immunogold labeling of AMPA and metabotropic receptors often is associated with tubulovesicular membranes of the ER, identified by the presence of BiP or calnexin (Fig. 10), indicating that this system could be a major route for the transport of dendritic proteins (Rubio and Wenthold, 1999a,b). The majority of long-distance organelle transport events in axons and dendrites are thought to be achieved by the active movements of microtubule-associated motor proteins, such as kinesins and cytoplasmic dyneins, along microtubule tracks (Hirokawa, 1998). Although characterization of organelle movement and associated proteins has been probed mostly in axons, there is evidence that a similar mechanism occurs in dendrites. This evidence includes: (1) the mixed orientation of 168

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microtubules in dendrites that support organelle transport (Baas et al., 1988; Overly et al., 1996); (2) the identification in dendrites, of motor proteins of the kinesin family, such as KIFC2 and KIF21B (Marszalek et al., 1999); and (3) the observation in vitro that organelles can reverse direction by changing motor activation or association, or switch to another microtubule (Brady et al., 1982; Smith and Forman, 1988). Myosins, which are involved in organelle transport in many systems, also are found in dendrites, and may play a role in dendritic protein movement (Mermall et al., 1998). In our analysis of the distribution of intracellular receptors in fusiform cells of the dorsal cochlear nucleus (Rubio and Wenthold, 1999a), we did not find evidence for a pool of receptors concentrated near the synapse (Fig. 5). Such a pool has been postulated as a receptor reserve that would allow a rapid insertion of additional receptors into the postsynaptic membrane. For example, long-term potentiation (LTP) has been suggested to involve the addition of AMPA receptors to the postsynaptic membrane, with a likely source of these receptors being an intracellular pool (Nayak et al., 1998; Morales and Goda, 1999; Shi et al., 1999). Since our results show a rather uniform distribution of receptors in the dendrite, recruitment of additional synaptic receptors would involve obtaining them from throughout the dendrite. If the addition of intracellular receptors is a component of LTP or other mechanisms involving rather rapid changes, the dendrite must utilize a mechanism to efficiently move these receptors. 3.3. FUNCTIONAL CONSIDERATIONS

3.3.1. Targeting mechanisms Differential distribution of glutamate receptors among two or more populations of synapses on a neuron implies that the cell utilizes some mechanisms to specifically target the receptors to the synapses (reviews of: Ehlers et al., 1996; Kirsch et al., 1996; Petralia et al., 1999b,c,d). Based on the distribution of mRNA, glutamate receptors appear to be synthesized predominantly in the neuronal cell body (Craig et al., 1993; Eshhar et al., 1993; Hunter et al., 1993; Laurie and Seeburg, 1994; Bahn and Wisden, 1997; see discussion in Gazzaley et al., 1997). Therefore, receptor expression at the postsynaptic plasma membrane requires an effective mechanism to selectively move receptors to their appropriate locations throughout the somatodendritic compartment. As noted above, receptors are transported somehow along the tubulovesicular network. In the dendrites, these receptors may rely on kinesins or cytoplasmic dyneins to affect transport of the membrane-bound receptors along microtubule pathways (Hirokawa, 1998). Final movement of the receptors through the synaptic spines presumably involves myosin motors traveling on actin pathways (Mermall et al., 1998; Naisbitt et al., 2000). Differential targeting of glutamate receptors to synapses may involve one of two processes: (1) receptors are transported along separate routes to different synaptic populations; or (2) receptors are transported indiscriminately throughout the somatodendritic compartment and are sorted only at the individual synapse. The first model, where receptors are transported along separate pathways, is supported by studies showing differential targeting of proteins to apical and basolateral membranes of polarized epithelial cells (Dotti and Simons, 1990; Perez-Velazquez and Angelides, 1993; Drubin and Nelson, 1996; Wozniak and Limbird, 1996). Separate pathways for glutamate receptors in neurons may be indicated by immunolabeling patterns in the cytoplasm. In the fusiform cell of the dorsal cochlear nucleus the synaptic labeling of glutamate receptors differs for apical and basal dendrites, as noted above. Interestingly, the cytoplasmic pools of 169

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,,,

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receptors show a corresponding distribution (Rubio and Wenthold, 1999a) (Figs. 5, 10 and 11). Thus, receptors (GluR2/3 immunolabeling) that are present at both apical and basal synaptic populations are equally abundant in apical and basal dendrite cytoplasm. In contrast, GluR4 and m G l u R l ~ , which are only present at the auditory nerve synapses on the basal dendrites, appear concentrated only in the cytoplasm of basal dendrites, being almost absent from apical dendrites. This indicates that a general targeting mechanisms exists to target receptors soon after their synthesis. At least in the case of targeting of GluR4 and m G l u R l ~ , the targeting mechanism may restrict movement of these receptors to the basal dendrites, which contain the target synapses. In the second model, receptors are transported indiscriminately throughout the somatodendritic compartment and are sorted only at the individual synapse. For example, some proteins are, transported throughout polarized epithelial cells and then are retained selectively at one pole (Wozniak and Limbird, 1996). Sorting mechanisms at the synapse may select which receptors can be inserted into the synapse (see below), or they may involve a selective attachment of glutamate receptors to associated proteins below the postsynaptic membrane (Fig. 7). The major group of these are: GRIP1, GRIP2, PICK1, A B E NSF and SAP-97 associated with AMPA receptors; PSD-95 associated with kainate receptors; PSD-93 associated with delta receptors; and PSD-95, SAP-102, and PSD-93 associated with N M D A receptors (Ziff, 1997; Kim and Huganir, 1999; Roche et al., 1999). Such anchoring mechanisms may vary according to the type of synapse or the functional state of the synapse. For example, based on the differential effects of detergent extraction on GluR1 clusters at synapses, Allison et al. (1998) suggest that AMPA receptors in cultured hippocampal neurons are anchored differently at two kinds of synapses, i.e., via GRIP protein in dendrite shaft synapses and via a weaker mechanism such as a spectrin 'corral' in dendrite spine synapses. Thus, the anchoring mechanism may effect differential distribution of the receptor. However, the necessity of anchoring mechanisms for receptor targeting has been questioned. Mutant mice that produce N M D A receptors lacking the binding site for anchoring proteins show significant functional disorders, yet still produce gateable, synaptically activated N M D A receptors, apparently independent

+_.__

Fig. 10. Double immunogold labeling with polyclonal antibodies for AMPA receptor subunits and BiP or calnexin in fusiform cell dendrites. (A, B) Double immunogold labeling with a polyclonal antibody for GluR2/3 (5-nm gold particles) and a monoclonal antibody for BiP (15 rim) in a basal proximal dendrite (A, BDI), and in a secondary apical dendrite (B, ADII). In both dendrites, membranes of the endoplasmic reticulum immunogold-labeled with BiP (15 nm) as well as GIuR2/3 (5 nm) (A, 1 and 3; B, 1). Some membranes that labeled only for BiP (arrows in A and B) and only for GluR2/3 (B, 2) are also observed. In (A), an auditory nerve terminal (AN) is seen making synaptic contact on a dendritic spine (1) on basal dendrites. The postsynaptic membrane (arrowheads) contains 5-nm gold particles specific for GluR2/3, and the cytoplasm of the spine (1)contains 15-rim gold particles specific for BiE (C, D) Proximal (BDI) and distal (BDII) basal dendrites, respectively, after double immunogold labeling with a polyclonal antibody specific for GluR4 (5 nm) and a monoclonal antibody for BiP (15 nm). In both dendrites, membranes of the endoplasmic reticulum showed immunogold labeling for BiP (15 rim) dispersed in the cytoplasm of the dendrite. Gold particles labeling GluR4 (5 nm; D, arrow) are associated with the same membranes labeled for BiP (arrowheads), but mostly are seen forming groups (C, arrows; D, 1 and 2) of particles associated with membranes that do not contain labeling for BiE (E) Double immunogold labeling with a polyclonal antibody specific for GluR2/3 (5 nm) and calnexin (15 nm) in a distal basal dendrite (BDII) of a fusiform cell. Colocalization of 5- and 15-nm gold particles is observed in a vesicle-like structure close to the plasma membrane adjacent to an auditory nerve synapse (AN). Gold particles labeling GluR2/3 are observed at the postsynaptic membrane of the auditory nerve (arrowheads) and in a smooth membrane in the cytoplasm (arrow). Insets show higher magnification of (A): 1-3. Scale bar, 0.25 ~m; insets, 50 rim; (E) 0.12 ~m. From Rubio and Wenthold (1999a). 171

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T

Fig. 11. Histograms showing the relative density of gold particles and SEM for GluR2/3, GluR4, and m G l u R l a (Ab-2) in different dendritic segments of fusiform cells (see Figs. 5 and 10). GluR2/3 immunolabeling is present in all dendritic segments, but with the highest level in both distal apical and basal dendritic segments (the difference between these two distal dendritic segments is not statistically significant, p > 0.01; the difference among the rest of the dendritic segments is statistically significant, p < 0.01). On the other hand, only basal dendrites show relatively high levels for GluR4 and mGluRl~. The labeling decreases toward the apical distal dendrites and is statistically significant (p < 0.01). The density of gold particles was compared for all the dendritic segments, and the difference of gold labeling was statistically significant for all cases (p < 0.01). As a control, we quantified the level of labeling in presynaptic areas adjacent to the dendrites of fusiform cells. The density (• of gold particles in presynaptic terminals was as follows: GluR2/3, 0.55 4- 0.16; GluR4, 0.92 4- 0.23; and mGluRla, Ab-1, 1.30 -+- 0.43; Ab-2, 1.16 + 0.27. ADIII - apical tertiary dendrite; ADII = apical secondary dendrite; ADI = apical primary dendrite; BDI = proximal primary basal dendrite; BDII --- distal secondary basal dendrite. From Rubio and Wenthold (1999a).

of any anchoring protein association (Sprengel et al., 1998). Such studies indicate that the anchoring proteins are not really anchors; rather they may be important as links between the receptors and transduction mechanisms that mediate the neuron's responses to activation of glutamate receptors (reviews by: Pawson and Scott, 1997; Craven and Bredt, 1998). In contrast, Moil et al. (1998) found evidence that similar mutant mice show deficits in the synaptic localization of NMDA receptors. Thus, the importance of anchoring mechanisms for receptor targeting is still unclear. 3.3.2. Insertion and removal of receptors at the synapse

The possible mechanisms for insertion and removal of glutamate receptors at synapses have been reviewed recently (Petralia et al., 1999d). Receptors may be incorporated into 172

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synapses either (1) by exocytotic insertion into the nonsynaptic membrane followed by lateral diffusion to the synapse (Baude et al., 1995), or (2) by direct incorporation from cytoplasmic tubulovesicular compartments into the postsynaptic membrane, presumably via an exocytotic process (Fig. 7). Evidence for both exocytosis at the sides of spines, and for connections of the spine reticulum to the postsynaptic density and membrane, have been described (Spacek and Harris, 1997; Petralia et al., 1999b,d). Presumably, glutamate receptors first traverse spans of the reticular network (see above; Rubio and Wenthold, 1999a,b; also general reviews by: Mironov et al., 1997; Nakata et al., 1998; Allan and Balch, 1999), and then either pass into vesicles and are exocytosed on the membrane of the dendrite shaft or dendrite spine (method 1), or are incorporated somehow directly into the postsynaptic membrane (method 2). Possibly there is more than one mechanism at a single synapse. For example, since the metabotropic receptors, mGluRl~ and mGluR5, are concentrated perisynaptically, it is possible that spines have different processes for inserting ionotropic and metabotropic glutamate receptors. Removal of receptors presumably involves some form of endocytosis, either at the postsynaptic membrane or on the spine or dendrite. Again, more than one method could occur; for example, muscarinic receptors may employ two or more mechanisms of endocytosis, including clathrin-coated pits and caveolae, depending on cell type and/or receptor subtype (Tolbert and Lameh, 1996; Feron et al., 1997; V6gler et al., 1998).

4. DISTRIBUTION IN GLIA Glutamate receptors are fairly common in glial cells (reviews by: Gallo and Russell, 1995; Steinh~iuser and Gallo, 1996). All four AMPA receptor subunits, GluR1-4, have been described in glia; at least in the gray matter of the cortex and hippocampus, GluR4 is the major glial AMPA receptor subunit (Jensen and Chiu, 1993; Conti et al., 1994; Gallo et al., 1994; Wenthold et al., 1996; Garcfa-Barcina and Matute, 1998). In the bovine corpus callosum, GluR1 is abundant in astrocytic end-feet and in the glial fibers surrounding the capillaries (Matute et al., 1994). GluR1 and GluR4 are abundant in Bergmann glia of the cerebellum (Petralia and Wenthold, 1992; Martin et al., 1993a; Sato et al., 1993a; Baude et al., 1994). Immunogold localization for GluR1 and GluR4 is seen in Bergmann glial processes that surround dendrites, dendritic spines and cell bodies of Purkinje cells (Rubio and Wenthold, 1999a). The kainate receptor subunits, GluR6, GluR7, KA1, and KA2 also are expressed to some extent in glial cells (Wisden and Seeburg, 1993; Gallo et al., 1994). Some evidence of glial labeling can be seen with delta 1/2 antibody (Mayat et al., 1995). Relatively little is known about NMDA receptors of glia (Uchihori and Puro, 1993; Petralia et al., 1994a). NR1 and NR2B mRNA have been reported in presumptive Bergmann glia (Luque and Richards, 1995), consistent with physiological studies (Mtiller et al., 1993). Immunolabeling for NR1 (Puro et al., 1996) and NR2A (Goebel et al., 1998) is seen in Mtiller cells, which are retinal radial glial cells similar to Bergmann glia. This is consistent with pharmacological evidence for NMDAreceptor-mediated effects of glutamate on glial cell proliferation and inhibition of potassium currents in the retina (Uchihori and Puro, 1993; Puro et al., 1996). In addition, GluR4, delta 1 and NR1 are found in glia in peripheral ganglia (Safieddine and Eybalin, 1992; Tachibana et al., 1994; Niedzielski and Wenthold, 1995; Safieddine and Wenthold, 1997) (Fig. 4). Numerous functions have been attributed to glial glutamate receptors on astrocytes, which may respond to glutamate released from synapses, and on oligodendrocytes, which may respond to glutamate released from axons (Steinhfiuser and Gallo, 1996). Since glial processes (from astrocytes and Bergmann glia) enwrap many kinds of excitatory synapses, 173

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the glial cell m a y l o c a l l y m o d u l a t e n e u r o t r a n s m i s s i o n (Schell et al., 1995). In addition, in r e s p o n s e to g l u t a m a t e , glia can (1) m o d i f y their ion c h a n n e l functions, p o s s i b l y to control the synaptic e n v i r o n m e n t (e.g., p o t a s s i u m h o m e o s t a s i s affecting n e u r o n a l excitability; Puro et al., 1996), (2) initiate c a l c i u m - d e p e n d e n t g e n e transcription p r o g r a m s , and (3) proliferate and differentiate (Steinh~iuser and Gallo, 1996).

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Targeting and anchoring of glutamate receptors and associated signaling molecules M. WYSZYNSKI AND M. SHENG

1. INTRODUCTION Glutamate acts upon multiple classes of receptors, but this chapter will focus on the wellstudied glutamate receptors that are concentrated in the postsynaptic membrane. Postsynaptic glutamate receptors can be classified as ionotropic receptors (iGluRs), which can be further divided into NMDA receptors, AMPA receptors, kainate receptors, and delta receptors; and metabotropic receptors (mGluRs). The group I mGluRs mGluRl~ and mGluR5, which are linked to phospholipase C (PLC) and phosphoinositide turnover, are predominant at postsynaptic sites. Both iGluRs and mGluRs have been studied intensively at the biophysical, pharmacological and electrophysiological levels, both in native preparations and in heterologous expression systems. In recent years, it has become apparent that glutamate receptors are specifically targeted to postsynaptic domains in neurons, indeed, even to subdomains within the postsynaptic specialization. At these specialized microdomains, glutamate receptors are associated with specific cytoplasmic proteins that link them to the cytoskeleton and to intracellular signal transduction pathways. Binding to specific intracellular proteins is likely to be important for immobilization and clustering of glutamate receptors, for their correct localization at postsynaptic sites, for their ability to transmit signals to appropriate cytoplasmic pathways, and for functional modulation of the receptors by kinases, phosphatases, and other regulatory proteins. Thus an intricate molecular machinery is involved in the subcellular targeting of glutamate receptors and in the assembly of receptor-associated protein complexes. Elucidation of these molecular mechanisms should reveal a great deal about the function and regulation of glutamate receptors in particular, and of excitatory synapses in general.

2. NMDA RECEPTORS

2.1. ASSOCIATION OF NMDA RECEPTORS WITH THE PSD Among the GluRs, NMDA receptors are biochemically the most tightly associated with the postsynaptic density (PSD), a morphological characteristic of excitatory synapses that is specialized for postsynaptic signal transduction (Kennedy, 1997; Ziff, 1997). Presumably, NMDA receptors are anchored in the PSD through specific protein-protein interactions mediated by the cytoplasmic domains of its constituent subunits. Via these biochemical interactions, NMDA receptors can be linked to the subsynaptic cytoskeleton and coupled to Handbook of Chemical Neuroanatomy, Vol. 18: Glutamate O.P. Ottersen and J. Storm-Mathisen, editors (~) 2000 Elsevier Science B.V. All rights reserved.

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_

i:f~ i,!~i ,

,

~.~;~ O i

Fig. 1. The NMDA receptor-PSD-95 complex. The C-terminus of NMDA receptor NR2 subunits binds to the first two PDZ domains of PSD-95. Major cytoplasmic components of the PSD-95 complex and their sites of binding are shown (see text for details). PSD-95 is shown multimerized via its N-terminal region (Hsueh et al., 1997), which is palmitoylated (Topinka and Bredt, 1998). Palmitoylation of N-terminal cysteines is important for targeting of PSD-95 to synaptic membranes (Craven et al., 1999). Individual PDZ domains are labeled 1, 2, 3. S -- SH3 domain; GK -- guanylate kinase-like domain.

postsynaptic signaling proteins (Fig. 1). Cytoskeletal interactions are functionally relevant because NMDA receptor activity is influenced by the actin cytoskeleton (Rosenmund and Westbrook, 1993; Paoletti and Ascher, 1994). The specificity of signal transduction is often determined by the nature of the molecular complex associated with the transmembrane receptor. This concept is likely to apply to NMDA receptor signaling because calcium influx through NMDA receptors stimulates specific intracellular events (e.g. synaptic plasticity, neurotoxicity, and transcriptional responses in the nucleus) that are not seen with other modes of calcium entry into the cell (Dingledine et al., 1999). Recent studies have uncovered many specific protein interactions mediated by the cytoplasmic tails of NMDA receptor subunits (Fig. 2). These findings are providing mechanistic insight into the synaptic targeting and signaling properties of NMDA receptors. 2.2. INTERACTIONS OF THE NR2 SUBUNIT: THE PSD-95 COMPLEX NMDA receptors are heteromeric (probably tetrameric) complexes composed of NR1 and NR2 subunits (Dingledine et al., 1999; Hollmann and Heinemann, 1994). There are four different NR2 subunits (NR2A-D), all of which have long cytoplasmic tails (up to 644 aa residues), the C-termini of which end in the conserved sequence -ESDV (NR2A, NR2B) or -ESEV (NR2C, NR2D). This short C-terminal peptide motif mediates binding to the PSD-95/SAP90 family of proteins, which are abundant core components of the PSD (Kornau et al., 1995, 1997; Niethammer et al., 1996; Sheng, 1996; O'Brien et al., 1998). PSD-95/SAP90 belongs to the MAGUK superfamily of proteins, which are characterized by the presence of PDZ domains, an SH3 domain and a guanylate kinase-like (GK) domain (Cho et al., 1992; Kistner et al., 1993). PDZ domains are modular protein domains of ~90 amino acids that are specialized for binding to C-terminal peptides in a sequence-specific 184

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((

Yotiao

X F-actin

Fig. 2. NMDA receptor interactions mediated independently of PSD-95. C1, C2 are alternatively spliced segments of the NR1 cytoplasmic tail. Black filled ovals represent actin-binding domains of 0~-actinin and spectrin. CaM = CaZ+/calmodulin; CaMKII = calmodulin-dependent kinase type II; PP1 = protein phosphatase 1; PKA = protein kinase A; NF-L = neurofilament-L.

fashion (Doyle et al., 1996; Cowburn, 1997; Ponting et al., 1997; Songyang et al., 1997). However, other modes of interaction are also possible with PDZ domains, including binding to internal sequences that fold into a 'beta-finger' (Hillier et al., 1999). PSD-95 has three PDZ domains in its N-terminal region; recognition of the ESDV C-terminal sequence of NR2 subunits is mediated by the first two PDZ domains (PDZ1 and PDZ2). Other members of the PSD-95 family in mammals include PSD-93/chapsyn-110 (Brenman et al., 1996b; Kim et al., 1996), SAP97/hDlg (Lue et al., 1994; Mtiller et al., 1995), and SAP102 (Miiller et al., 1996). All the family members except SAP97 (which is reported to be predominantly presynaptic and axonal) (Mtiller et al., 1995) appear to be components of the PSD and to be associated with NMDA receptors in synapses. While few dispute the existence of this interaction in vivo, the functional significance of PSD-95 binding to NMDA receptors remains incompletely understood. PSD-95 may be involved in the synaptic targeting of NMDA receptors, in the coupling of NMDA receptors to signaling proteins, or in the anchoring of NMDA receptors to the postsynaptic cytoskeleton, or in a combination of these functions. 2.3. SYNAPTIC TARGETING BY PSD-95 An early hypothesis based on the co-localization of PSD-95 and NMDA receptors in the PSD is that NR2 binding to PSD-95 is important for the postsynaptic localization of NMDA receptors. This idea was supported by genetic experiments in drosophila on the fly homolog of PSD-95, Discs large (Dlg) (Woods and Bryant, 1991). Dlg is concentrated in the NMJ of drosophila, a glutamatergic synapse (Guan et al., 1996), where it co-localizes with the 185

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Shaker K + channel and the Fasciclin II (FasII) cell adhesion molecule, two transmembrane proteins that bind directly to the PDZ domains of Dlg. dlg mutants show loss of the normal synaptic localization of Shaker and FasII (Tejedor et al., 1997; Thomas et al., 1997; Zito et al., 1997). Moreover, the C-termini of FasII and Shaker (containing the PDZ-binding motifs) were sufficient to confer synaptic targeting on a heterologous protein in wild-type but not in dlg mutant flies (Zito et al., 1997). Taken together, these genetic studies indicated that Dlg is important in vivo for synaptic localization of its membrane protein binding partners. Drosophila glutamate receptors have not been shown to bind to Dlg, and it remains unclear how ionotropic glutamate receptors are targeted to postsynaptic sites in the drosophila NMJ. Thus a direct homology with mammalian NMDA receptors and PSD-95 is not available. In Caenorhabditis elegans (C. elegans), genetic experiments implicate another PDZ-containing protein (LIN-10, which is not a MAGUK protein) in synaptic localization of ionotropic glutamate receptors (Rongo et al., 1998). By extrapolation from genetic studies in drosophila and C. elegans, it was natural to speculate that the PSD-95 family of proteins in mammals would be involved in the targeting of NMDA receptors to the postsynaptic specialization. However, direct evidence for this has not been forthcoming. A 'knockout' of the PSD-95 gene in mice did not cause a detectable defect in synaptic localization of NMDA receptors, although downstream signaling functions of NMDA receptors were apparently altered (Migaud et al., 1998). (However, it could be argued that close relatives of PSD-95 could compensate for loss of PSD-95.) Dominant interfering approaches with peptides that compete for PDZ binding also argued that PSD-95 and its relatives are not essential for normal targeting of NMDA receptors (Passafaro et al., 1999). Similarly, mice with targeted deletions of the cytoplasmic tails of NR2A, NR2B, and NR2C also had apparently normal synaptic localization of the mutant NMDA receptors (Sprengel et al., 1998). In contrast, another study did find a significant loss of synaptic localization of NR2B in mice expressing a 'tail-less' NR2B (Mori et al., 1998). However, tLaese results are complicated by the deleterious effects of this mutation on brain development and organismal survival (Mori et al., 1998). In general, it appears that the NR2 interaction with PSD-95 family proteins is not absolutely essential for synaptic targeting of NMDA receptors. This is most likely because other (redundant) mechanisms exist for the proper localization of NMDA receptors, e.g. via interactions with NR1 (see below). In this context, it would be interesting to study the synaptic targeting of PSD-95-interacting proteins other than NMDA receptors to see whether their distribution is affected by loss of PSD-95 function. PSD-95 itself has multiple determinants within its primary structure that are required to target it to postsynaptic sites (Arnold and Clapham, 1999; Craven et al., 1999). 2.4. ASSEMBLY OF A SIGNALING COMPLEX BY PSD-95 In contrast to the equivocal results with respect to PSD-95's role in synaptic targeting of NMDA receptors, genetic experiments in mice provide convincing evidence that the NR2-PSD-95 interaction is critical for intracellular signaling by the NMDA receptor. The PSD-95 knockout mice showed dramatic changes in NMDA receptor-dependent synaptic plasticity, shifting the threshold between LTP and LTD, greatly enhancing LTP magnitude, and disrupting spatial learning (Migaud et al., 1998). Rather surprisingly, the mutant phenotype suggests that PSD-95 normally has a constraining influence on LTR perhaps by linking NMDA receptors to negative downstream regulators of synaptic transmission, such as protein phosphatases. Further evidence that NR2 interactions with cytoplasmic proteins are important for NMDA receptor signaling comes from mouse mutants that have targeted deletions of 186

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the cytoplasmic tails of NR2A, NR2B, and NR2C (Sprengel et al., 1998). These mutations essentially phenocopied the deletion of the entire respective genes (in the case of NR2A and NR2C, without obviously affecting NMDA receptor expression or channel activity). It should be noted that these targeted mutations deleted the entire cytoplasmic tails (~400-600 amino acids) of the NR2 subunits, while only the last few amino acids are involved in binding to PSD-95. Thus these 'tail-deletion' phenotypes may be due to loss of functions other than PSD-95 binding. One interaction that may be affected is with calmodulin-dependent protein kinase II (CaMKII), which has been reported to bind to proximal regions of NR2B cytoplasmic tail (Strack and Colbran, 1998; Leonard et al., 1999). The functional role of a direct interaction between NMDA receptor subunits and CaMKII is unclear, though NMDA receptors are substrates for CaMKII (Omkumar et al., 1996). The cytoplasmic tail of NR2 subunits are up to ~600 amino acids in length, so other protein interactions and other functions almost certainly reside in this extended region (see below). Why is PSD-95 important for the signaling functions of NMDA receptors? Probably because PSD-95 binds to a variety of cytoplasmic proteins that are involved in downstream signaling of NMDA receptors (see Fig. 1; and reviewed in Craven and Bredt, 1998). Accumulating evidence indicates that PSD-95 functions as a scaffold for assembling a specific protein complex associated with NMDA receptors. For example, neuronal nitric oxide synthase (nNOS; which itself contains a PDZ domain) has been shown to bind to PDZ2 of PSD-95 via a PDZ-PDZ interaction (Brenman et al., 1996a). nNOS is a calcium/calmodulin-regulated enzyme that is selectively activated by calcium influx through NMDA receptors (as opposed to calcium entry through voltage-gated calcium channels). This specific coupling can be neatly explained by the physical approximation of NMDA receptor and nNOS through their mutual binding to PSD-95. Significantly, antisense knockdown of PSD-95 inhibits nNOS activation in response to NMDA receptor stimulation and suppresses NMDA receptor-mediated excitotoxicity in cultured neurons (Sattler et al., 1999). A variety of signaling proteins have now been identified that interact directly with PSD-95 and that are presumably therefore associated indirectly with NMDA receptors. These include regulators or effectors of Ras and Rho GTPases. SynGAR a GTPase activating protein for Ras, has a C-terminus that interacts with all three PDZ domains of PSD-95 (Chen et al., 1998; Kim et al., 1998). SynGAP is a PSD protein whose association with PSD-95 positions it close to NMDA receptors perhaps SynGAP functions to inactivate Ras that is activated locally by NMDA receptor stimulation (Yun et al., 1998). SynGAP may also be involved in Ras modulation following other modes of Ras activation, such as by postsynaptic receptor tyrosine kinases. The functional role of SynGAP in synaptic function and plasticity remains to be clarified; it is a large protein that may have additional activities unrelated to its RasGAP domain. Citron, a putative effector for Rho, can also bind PSD-95, specifically via PDZ3 (Furuyashiki et al., 1999; Zhang et al., 1999). Since Rho-type GTPases are involved in regulation of the cytoskeleton, it is speculated that activity-dependent modulation of postsynaptic actin might be mediated via a cascade involving NMDA receptors, Rho/Rac, PSD-95, and Citron. However, in the hippocampus, Citron is concentrated only in glutamatergic synapses of inhibitory neurons (Zhang et al., 1999). This finding illustrates the principle that distinct protein complexes can be assembled around PSD-95 (and hence NMDA receptors) in different neuronal cell types. An emerging theme in cell biology is that protein kinases are often targeted to their substrates by association with specific anchoring proteins. Non-receptor tyrosine kinases of the Src family have been implicated in NMDA receptor modulation (Salter, 1998), and recent evidence suggests that Src family kinases may be components of the NMDA 187

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receptor-associated protein complex. Fyn is a major tyrosine kinase that phosphorylates the NR2A subunit in vivo. Association of Fyn with NMDA receptors appears to be mediated by binding of the SH2 domain of Fyn to PDZ3 of PSD-95 (Tezuka et al., 1999). Indeed, coexpression of PSD-95 enhances the phosphorylation of NR2A by Fyn in heterologous cells (Tezuka et al., 1999), suggesting that PSD-95 targets Fyn to the NMDA receptor by mediating a ternary complex. Other tyrosine kinases of this family (Src, Yes, and Lyn) can also be co-immunoprecipitated with NMDA receptors (Yu et al., 1997; Tezuka et al., 1999), although binding to PSD-95 has yet to be shown. Thus PSD-95 family proteins may play a role in bringing Src-like tyrosine kinases to the NMDA receptor complex. The guanylate kinase-like (GK) domain of PSD-95 family proteins shows sequence similarity to the enzyme guanylate kinase, but no catalytic activity has been found for this domain (Kuhlendahl et al., 1998). Instead, the GK domain appears to act as another site for protein-protein interaction that can link NMDA receptors indirectly to cytoplasmic signaling proteins. The GK domain binds to an abundant family of proteins in the PSD, termed GKAP/SAPAP/DAP, whose function is unclear (Kim et al., 1997; Naisbitt et al., 1997; Satoh et al., 1997; Takeuchi et al., 1997). In addition, the GK domain binds to BEGAIN, a novel protein of unknown function (Deguchi et al., 1998); SpanGAP, a putative GTPase activating protein for the small GTPase Rap (D. Pak and M. Sheng, unpublished observations); and MAP1A, a microtubule-binding protein whose binding to the GK domain is stimulated by occupancy of the neighboring PDZ domains (Brenman et al., 1998). It is unclear what these GK-binding proteins are doing for NMDA receptor function or signaling. Some of these may serve 'structural' roles to link NMDA receptors to other proteins. For instance, GKAP has recently been shown to bind to Shank, a scaffold protein that in turn binds to Homer (see below) (Naisbitt et al., 1999; Tu et al., 1999). This chain of protein-protein interactions could couple NMDA receptors to intracellular calcium stores, since Homer interacts with inositol 1,4,5-trisphosphate receptors (IP3R) and appears to be generally involved in excitation-calcium coupling (see Section 5). Further studies are needed to characterize the functional significance of the PSD-95-based complex in NMDA receptor signaling and modulation. It should be borne in mind that PSD-95 might organize other membrane receptors in addition to NMDA receptors (such as adhesion receptors or receptor tyrosine kinases), and thus PSD-95-associated proteins may serve NMDA receptor-independent signaling functions. Like other MAGUKs, PSD-95 family proteins contain an SH3 domain, a well-known protein-binding module (Pawson, 1995). The only binding partner that has been identified for the SH3 domain of PSD-95 is a kainate receptor subunit (Garcia et al., 1998) (see below). In addition, an intriguing intramolecular interaction has been identified between the SH3 domain and the GK domain of PSD-95 family proteins (Kim and Sheng, 1999; McGee and Bredt, 1999). The significance of this SH3-GK intramolecular interaction is unknown. The PSD-95-based protein complex linked to NMDA receptors will probably continue to grow in size and complexity in the coming years. So far the rate of discovery of PSD-95-interacting proteins has greatly outpaced our understanding of the functional significance of these proteins. 2.5. ANCHORING TO THE CYTOSKELETON VIA PSD-95 Anchoring to the subsynaptic cytoskeleton can be considered a final step in the process of postsynaptic targeting of glutamate receptors. By binding to cytoskeletal elements, a scaffold protein such as PSD-95 can indirectly connect NMDA receptors to the cytoskeleton. 188

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Members of the PSD-95 family of proteins have been shown to bind in vitro to band 4.1, an actin/spectrin-binding protein (Lue et al., 1994, 1996; Marfatia et al., 1996). Such an interaction has the potential to link NMDA receptors indirectly to F-actin, which is the predominant cytoskeleton in dendritic spines. Whether band 4.1 or other members of the ezrin-radixin-moesin (ERM) family of actin-binding proteins play a role in postsynaptic anchoring of NMDA receptors and PSD-95 is unknown. PSD-95 also interacts with microtubule-associated proteins. This is somewhat surprising, since microtubules are generally thought to be sparse or absent from dendritic spines. Nevertheless, tubulin is present in PSD preparations and microtubule-associated proteins such as MAP2 have been immunocytochemically localized at synapses (Kelly and Cotman, 1978; Caceres et al., 1984; Walsh and Kuruc, 1992). The third PDZ domain of PSD-95 binds to CRIPT, a small polypeptide which binds directly to microtubules (Niethammer et al., 1998; Passafaro et al., 1999). The interaction of PSD-95 family proteins with microtubule-binding proteins such as CRIPT or MAP1A (see above) may link NMDA receptors to a postsynaptic tubulin-based cytoskeleton. It is controversial whether tubulin contributes to the cytoskeletal organization of the PSD in dendritic spines (Harris and Kater, 1994; Lai et al., 1998). However, microtubule anchoring may be relevant for the minor fraction of excitatory synapses that are made on to microtubule-rich dendritic shafts (such as the aspiny excitatory synapses of inhibitory interneurons). Alternatively, microtubule interactions may be more related to trafficking of PSD-95 from the cell body than to cytoskeletal anchoring at synapses. 2.6. INTERACTIONS OF THE NR1 SUBUNIT NMDA receptors contain the essential NR1 subunit in addition to the NR2 subunits that bind to PSD-95 family proteins. The cytoplasmic tail of NR1 undergoes considerable alternative splicing (Hollmann et al., 1993). Although it does not bind to PSD-95, the C-terminal cytoplasmic tail of NR1 does interact with several other cytoplasmic proteins (Fig. 2). Like NR2-PSD-95 interactions, these NRl-mediated interactions may play a role in synaptic targeting of NMDA receptors and NMDA receptor-associated signaling proteins. ~-Actinin, an actin-binding protein of the spectrin superfamily, interacts with the membrane proximal segment (termed CO) of NRI's cytoplasmic tail that is common to all splice variants (Wyszynski et al., 1997, 1998a). Since ~-actinin is enriched in the PSD, its interaction with NR1 may contribute to NMDA receptor-cytoskeletal anchoring at postsynaptic sites. CaZ+/calmodulin binds to two distinct sites in the NR1 tail, to the CO segment (Ka ~ 80 nM) and to the C1 segment (Kd -- 3.7 nM), the latter segment being encoded by the differentially spliced exon 22 of the gene (Ehlers et al., 1996). The binding of CaZ+/calmodulin inhibits NMDA receptor opening and reduces mean channel open time (Ehlers et al., 1996). The calmodulin and ~-actinin binding sites overlap in CO, and these proteins compete in vitro for binding to NR1 (Wyszynski et al., 1997). This competition between calmodulin and ~-actinin appears to be involved in calcium-dependent inactivation of NMDA receptors, with inactivation occurring by the competitive displacement of ~-actinin from NR1 by CaZ+/calmodulin (Zhang et al., 1998; Krupp et al., 1999). The CO segment of the NR1 cytoplasmic tail is required for the calcium-dependent inactivation of NMDA receptors, and this part of the NR1 tail may be directly involved in channel gating. Thus if the actinin-NR1 interaction is involved in cytoskeletal anchoring of NMDA receptors, it is intimately tied to the gating of NMDA receptors. The C1 exon segment of the NR1 tail is not required for calcium-dependent inactivation of NMDA receptors despite binding calmodulin, but it does contain several protein kinase C 189

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phosphorylation sites that play a role in the clustering of NR1, at least when overexpressed in heterologous cells (Ehlers et al., 1995). Two proteins, yotiao (Lin et al., 1998) and neurofilament NF-L (Ehlers et al., 1998), have been found to interact specifically with splice variants of NR1 containing the C1 exon (Fig. 2). Yotiao has recently been identified as an A-kinase anchoring protein (or AKAP) that binds to both protein kinase A (PKA) and protein phosphatase 1 (PP1). Yotiao may function in synapses to organize a serine/threonine kinase-phosphatase complex closely linked to NMDA receptors, thus facilitating bidirectional NMDA receptor modulation by these enzymes (Westphal et al., 1999). Spectrin, a well-known actin-binding protein, is reported to bind to the cytoplasmic domains of NR1; it also has affinity for NR2A and NR2B (Wechsler and Teichberg, 1998). The spectrin binding site in NR2B is distinct from the ~-actinin and PSD-95 binding regions. A specific form of brain spectrin is abundant in the PSD, and may thus offer another mode for attaching NMDA receptors to the postsynaptic actin cytoskeleton. Spectrin interaction with NR2B is sensitive to tyrosine phosphorylation and calcium, whereas the binding of spectrin to NR1 is inhibited by PKC/PKA phosphorylation and calmodulin (Wechsler and Teichberg, 1998). These findings suggest possible mechanisms for activity-dependent regulation of NMDA receptor anchoring to the cytoskeleton. In this context, it is worth noting that the synaptic localization of NMDA receptors in cultured neurons can be enhanced by NMDA receptor antagonists, although the mechanism for this effect remains unknown (Rao and Craig, 1997). 2.7. OTHER INTERACTIONS OF NMDA RECEPTORS The list of proteins that can interact with the cytoplasmic tail of NMDA receptor subunits is growing. Here we discuss a few examples that were not mentioned in earlier sections. S-SCAM, a protein with an N-terminal GK domain followed by two WW motifs and five PDZ domains, has been shown to bind to NR2 subunits with its fifth PDZ domain (Hirao et al., 1998). S-SCAM belongs to a family of proteins that includes AIP1 and MAGI, which are distantly related to the MAGUK proteins. MALS, a mammalian homolog of LIN-7, can also bind to NR2 subunits via a PDZ-C-terminus interaction (Jo et al., 1999). The significance of these interactions for NMDA receptor function in vivo remains to be determined. In conclusion, it seems clear that NMDA receptors interact with a multitude of intracellular proteins, either directly or indirectly via scaffold proteins like PSD-95. Undoubtedly, there are many protein interactions involving NMDA receptors that remain to be uncovered. These interactions are likely to contribute to the cytoskeletal anchoring of NMDA receptors in the PSD, and to the coupling of NMDA receptors to intracellular signaling pathways.

3. AMPA RECEPTORS

3.1. SYNAPTIC TARGETING OF AMPA RECEPTORS Like NMDA receptors, AMPA receptors are also typically concentrated at postsynaptic sites of excitatory synapses. Recent evidence suggests, however, that the targeting of AMPA receptors to synapses is much more heterogeneous than that of NMDA receptors. Many excitatory synapses contain NMDA receptors but not AMPA receptors, especially early in development, and the content of AMPA receptors in AMPA receptor-positive synapses is quite variable (Nusser et al., 1998; Petralia et al., 1999; Takumi et al., 1999). Such 'morphologically silent 190

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G ....

a-kipdns

EphA7/B2 EphrinB

F/g. 3. The AMPA receptor-associated complex. AMPA receptors are shown as heteromers of GluR1 and GluR2. PICK-1 is depicted as a dimer; via interaction of its coiled-coil domains. NSF/SNAP binds to GluR2 cytoplasmic tail in an ATP-dependent manner. PDZ6 of GRIP is the binding site for ~-liprins and Eph receptors and their ligands, but binding partners for most of the PDZ domains of GRIP remain to be identified.

synapses' are also found in cultured neurons, where the synaptic expression of surface AMPA receptors can be altered by blocking NMDA or AMPA receptors and by activity (Carroll et al., 1999; Liao et al., 1999). The regulated targeting of AMPA receptors to postsynaptic sites may be involved in controlling synaptic efficacy, according to the 'silent synapse' hypothesis (Malenka and Nicoll, 1997). Thus the mechanisms for synaptic targeting of AMPA receptors are probably more complicated than for NMDA receptors and more immediately relevant to synaptic plasticity. AMPA receptors are typically composed of heteromeric combinations of GluR1-4 subunits (Hollmann and Heinemann, 1994; Dingledine et al., 1999), whose membrane topology is similar to that of NMDA receptor subunits. In analogous fashion, the C-terminal cytoplasmic tails of AMPA receptor subunits also interact with intracellular proteins (Fig. 3). Despite coexisting at the same excitatory synapses, however, AMPA receptors bind to a more distinct set of cytoplasmic proteins than NMDA receptors, presumably reflecting the differential regulation of these receptor channels. Most AMPA receptor binding proteins have been identified via interaction with the GluR2/3 subunits (Fig. 3). As with NMDA receptors, many AMPA receptor interactions are mediated by the binding of subunit C-termini to specific PDZ-containing scaffold proteins. 3.2. INTERACTIONS WITH PDZ PROTEINS GluR2 and GluR3 subunits share a common C-terminal sequence (-SVKI) that interacts with the fifth PDZ domain of GRIP (now termed GRIP1), a protein containing seven PDZs and no other recognizable domains (Dong et al., 1997; Wyszynski et al., 1998b). A protein with six PDZ domains (AMPA receptor binding protein or ABP) was also isolated by its binding 191

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to GluR2/3 (Srivastava et al., 1998). ABP appears to be a splice variant of a GRIP-related protein (also called GRIP2) that contains seven PDZs; ABP lacks the N-terminus and PDZ7 of GRIP2 (Bruckner et al., 1999; Dong et al., 1999; Wyszynski et al., 1999). Although a large fraction of GluR2/3 appears to be biochemically associated with GRIP in vivo (Wyszynski et al., 1999), the function of the GluR2/3-GRIP interaction is still unclear. At the subcellular level, GRIP is present in synapses in the brain but not as specifically as is PSD-95. Indeed, GRIP is rather weakly associated with synapses in culture, and its expression in the brain pre-dates AMPA receptors during development (Dong et al., 1999; Wyszynski et al., 1999). Thus GRIP almost certainly is involved in functions in addition to AMPA receptor anchoring in synapses. GRIP also differs from PSD-95 in being relatively abundant in intracellular compartments in dendrites and cell bodies of neurons, suggesting that GRIP may be more important for trafficking than for synaptic anchoring of AMPA receptors (Dong et al., 1999; Wyszynski et al., 1999). The fact that overexpression of the C-terminal tail of GluR2 in neurons inhibits synaptic clustering of AMPA receptors (Dong et al., 1997) is consistent with either an anchoring or trafficking role for GRIE Containing seven or six PDZ domains, respectively, GRIP and ABP have the capacity to assemble a large protein complex around AMPA receptors (Fig. 3). GRIP has been shown to bind to EphB2 and EphA7, members of the large family of Eph receptor tyrosine kinases, and to the EphrinB ligands for Eph receptors (Torres et al., 1998; Bruckner et al., 1999). Eph receptor-ephrin interactions are involved in axon guidance, cell migration, and establishment of tissue boundaries (Flanagan and Vanderhaeghen, 1998). Liprins, proteins that bind to the LAR family of receptor tyrosine phosphatases (Serra-Pag~s et al., 1998), also bind to GRIP, utilizing PDZ6 (M. Wyszynski, M. Sheng, unpublished observations). LAR tyrosine phosphatases are involved in axon guidance during neural development (Van Vactor, 1998), but appear to concentrate in synapses in mature neurons as do Eph receptors and ephrins (Torres et al., 1998). How GRIP-mediated interactions with Eph receptors and liprins are relevant to AMPA receptors is unclear at present, especially given that GRIP probably has functions unrelated to AMPA receptors. In addition to GRIP/ABE the C-terminal sequence of GluR2/3 mediates binding to PICK1 (Xia et al., 1999), another PDZ-containing protein previously shown to bind protein kinase C (PKC) (Staudinger et al., 1995). Thus, like NMDA receptors, AMPA receptor subunits have specific affinity for more than one PDZ domain protein. PICK1 co-localizes with GluR2 in synapses and is capable of clustering GluR2 in heterologous cells (Xia et al., 1999), perhaps via coiled-coil dimerization of PICK1. Since PKC~ is enriched in synapses, the possibility exists that PICK1 may recruit PKC to AMPA receptors, although this has yet to be demonstrated. The relative importance of PICK1 and GRIP/ABP in AMPA receptor anchoring/trafficking in vivo remains to be worked out. The GluR1 subunit of AMPA receptors does not bind to GRIP, ABE Or PICK1, but its C-terminus has been recently shown to associate with SAP97, a member of the PSD-95 family of MAGUKs (Leonard et al., 1998). Since SAP97 is reported to be predominantly presynaptic (MUller et al., 1995), however, the physiological significance of this interaction is uncertain. 3.3. INTERACTIONS WITH NSF AND SIGNALING PROTEINS As with NMDA receptors, C-terminal-PDZ interactions are not the only mechanism for linking AMPA receptors to intracellular proteins. A surprising finding (reported independently by three different research groups) was that GluR2 binds to NSF, an ATPase required for the vesicle fusion cycle (Nishimune et al., 1998; Osten et al., 1998; Song et al., 1998). NSF 192

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binding is mediated by a membrane proximal segment of GluR2's cytoplasmic tail, rather than the C-terminus that binds to GRIP or PICK1 (Fig. 3). Further, Osten et al. (1998) have shown that GluR2-NSF interacts with ~- and [~-soluble NSF attachment proteins (SNAPs), and that the assembly of the GluR2-NSF-SNAP complex is reversible by ATP hydrolysis. The functional significance of these interactions is not understood, but the obvious speculations are that NSF is involved in the vesicle trafficking or chaperoning of AMPA receptors (reviewed in Lin and Sheng, 1998). Additionally, AMPA receptors have been shown to be associated with the Src-related non-receptor tyrosine kinase Lyn, which co-immunoprecipitates with GluR2/3 from cerebellar extracts (Hayashi et al., 1999). This association requires the SH3 domain of Lyn and a membrane proximal 20 amino acid region of the GluR2 C-terminal tail (just upstream of the NSF binding site); however, a direct interaction between the two proteins has yet to be shown. Lyn is activated by AMPA receptor stimulation and is required for AMPA receptor-mediated stimulation of MAP kinase and BDNF gene expression, but this signaling is unusual in that it appears to be independent of ion flux by the AMPA receptor. Instead, the authors propose that the signal is transduced by conformational changes in the receptor upon binding of AMPA (Hayashi et al., 1999). This mechanism is reminiscent of the activation of a heterotrimeric G-protein (Gi) by AMPA receptors in cortical neurons (Wang et al., 1997), which also appears to be independent of GluR channel function.

4. KAINATE RECEPTORS AND 8 RECEPTORS

Kainate receptors represent a third class of glutamate-gated ion channel, and are made up of subunits (GluR5-7, KA1 and KA2) that are homologous to AMPA receptor subunits. The cytoplasmic domains of GluR6 and KA2 have been shown to bind to the PDZ1 domain and to the SH3 and GK domains of PSD-95, respectively (Garcia et al., 1998). Coexpression with PSD-95 can alter the desensitization properties of kainate receptors in heterologous expression systems. Another member of the ionotropic glutamate receptor superfamily is GluR~, distantly related (~25% identity) to NMDA and AMPA/kainate receptors. GluR~2, the most studied member of this family, is expressed specifically in cerebellar Purkinje cells. Gain-of-function mutations in the GluR82 gene underlie the phenotype of Lurcher mice (Zuo et al., 1997), while targeted gene disruption demonstrates a requirement for GluR32 in synapse function and development in cerebellum (Kashiwabuchi et al., 1995). GluR~2 binds to PSD-93/chapsyn-110 in vitro and co-localizes with PSD-93 in parallel-fiber Purkinje cell synapses in vivo, suggesting a possible role for PSD-93 in anchoring of GluR~2-containing glutamate receptors (Roche et al., 1999).

5. METABOTROPIC GLUTAMATE RECEPTORS

Glutamate acts on G-protein-coupled metabotropic receptors in addition to the ionotropic receptors discussed above. Metabotropic glutamate receptors (mGluRs) are divided into three classes based on G-protein coupling and pharmacology. Group 1 (mGluR1 and mGluR5) are predominantly postsynaptic and activate phospholipase C (PLC) and intracellular calcium release, whereas group 2 (mGluR2 and mGluR3) and group 3 (mGluR4, 6, 7, 8) receptors function at both pre- and postsynaptic sites and negatively couple adenylyl cyclase. Recent 193

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NMDA receptors and mGluRs are shown in the center, and around the periphery of the synapse, respectively. IP3 receptors (IP3R) are present in the smooth endoplasmic reticulum (SER), an intracellular store for Ca 2+. Monovalent Homer, Homerla (Hla), is an immediate early gene product that competes with the constitutively expressed multivalent Homer (H) for mGluR binding, thereby uncoupling mGluR from IP3R. mGluR, group 1 metabotropic glutamate receptor; SHK = Shank; G = GKAP; P - PSD-95.

evidence suggests that differential subcellular targeting among the mGluR family is probably determined by sequences in their cytoplasmic C-terminal tails (Stowell and Craig, 1999). This review focuses on group 1 mGluRs because their subcellular localization is best characterized. Although mGluR1 and mGluR5 interact with G-proteins like other 7-transmembrane receptors, we discuss only the more specific protein interactions that may be involved in targeting and signaling of these postsynaptic mGluRs. Unlike NMDA receptors and AMPA receptors, which are distributed across the PSD, group 1 mGluRs are concentrated in a ring around the periphery of the PSD (Nusser et al., 1994; Lujfin et al., 1997; Takumi et al., 1999). This segregation of ionotropic and metabotropic receptors at the subsynaptic level probably depends on differential interactions of these membrane proteins with cytoplasmic proteins. Brakeman et al. (1997) uncovered a specific interaction between mGluRl~ (a splice variant of mGluR1), mGluR5 and the cytoplasmic protein Homer (Fig. 4). Although the interaction was originally described as occurring between the C-terminus of m G l u R l ~ / 5 and a PDZ-like domain of Homer, subsequent analysis has revealed that the binding occurs between the EVH domain of Homer and an internal sequence motif (PPXXF) in the cytoplasmic tail of mGluR1/5 (Tu et al., 1998; Xiao et al., 1998). The originally identified Homer gene (now termed Homerla) was an immediate early gene whose mRNA was induced by synaptic activity (Brakeman et al., 1997). Subsequently, a family of Homer proteins was described containing a coiled-coil domain that mediates self-association (Kato et al., 1998; Xiao et al., 1998). These 'CC-Homers' can multimerize to form multivalent complexes that could crosslink multiple m G l u R l ~ / 5 molecules, or link mGluR10~/5 to other proteins containing the PPXXF Homer-binding motif (Xiao et al., 1998). In contrast, Homerla, which lacks the coiled-coil domain, cannot multimerize; instead, it behaves as a natural dominant negative to disrupt CC-Homer-mediated protein complexes (Xiao et al., 1998). Several other proteins have been noted to contain the PPXXF Homerbinding consensus, of which the most pertinent is the IP3 receptor (IP3R), a downstream effector in the m G l u R 1 / 5 - P L C signaling pathway, mGluR10~ can be co-immunoprecipitated as a complex with Homer and IP3R from rat cerebellum (Tu et al., 1999), consistent with a biochemical linkage between group 1 mGluRs and the IP3R. More importantly, overexpression of the interfering Homerla in Purkinje neurons delayed and inhibited mGluR-evoked 194

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intracellular calcium release (Tu et al., 1998). These findings argue that the physical tethering of mGluRl~/5 to IP3R via CC-Homer is functionally important for postsynaptic calcium responses to mGluR stimulation. These findings have led to the model that Homer brings IP3R into close proximity of the group 1 mGluRs (Fig. 4), thereby allowing for more efficient coupling between these proteins. The idea is that stimulation of mGluRl~/5 leads to highly localized production of IP3 at postsynaptic sites, such that the IP3R has to be in close vicinity. This is therefore an example of a signaling microdomain based on a protein complex (which contains at least mGluRl~/5, Homer, and IP3R). IP3Rs are associated with the smooth endoplasmic reticulum (SER), a calcium store found in dendritic spines that often approaches close to the postsynaptic specialization. So the morphological basis for a close interaction between postsynaptic receptors and intracellular calcium compartments exists in dendritic spines (Fig. 4). In this context, it is of interest that ryanodine receptors also contain the PPXXF consensus for Homer binding. Homer also binds to Shank, which contains a PPXXF motif in its proline-rich domain (Tu et al., 1999). Since Shank is a component of the NMDA receptor complex via binding to GKAP (Naisbitt et al., 1999), the Homer-Shank interaction potentially links the group 1 mGluRs to the NMDA receptor and its associated proteins (Fig. 4). In addition, the group 1 mGluRs may interact directly with Shank. The cytoplasmic tail of mGluR5 ends with a sequence (-SSSL) reminiscent of the terminal SXV consensus that binds to the PDZ domains of PSD-95. This C-terminal sequence is reported to bind to the PDZ domain of Shank, which preferentially recognizes the terminal T/SXL motif (Naisbitt et al., 1999; Tu et al., 1999). Thus mGluRla/5, Homer and Shank may form a 'triangular' complex with each other. Is the binding to Homer and/or Shank important for determining the perisynaptic location of group 1 mGluRs? This question has not been answered directly. One argument against such a targeting function is that although Homer and Shank are enriched in synapses, they are found throughout the PSD, in contrast to mGluR1/5, which are arranged around the periphery of the PSD. Thus the specific subsynaptic segregation of group 1 mGluRs cannot be explained simply by binding to Homer and Shank. However, Homer and Shank could contribute to the anchorage of mGluRs at postsynaptic sites.

6. C O N C L U D I N G C O M M E N T S : C O M P A R I N G GLUTAMATE RECEPTORS As detailed above, a dauntingly complicated picture has emerged of the interactions of glutamate receptors with cytoplasmic proteins. This seems particularly true of the NMDA receptors, which play diverse roles in postsynaptic signaling as a result of their calcium permeability. NMDA receptors utilize both NR1 and NR2 subunits to participate in multiple specific sets of interactions with cytoplasmic proteins. These NMDA receptor-interacting proteins may have direct effects on receptor-channel activity (such as ~-actinin and calmodulin) or they may function as adaptor/scaffold proteins (like PSD-95) that connect the receptor to a more complex network of postsynaptic molecules. NMDA receptors do not associate with microtubules or actin directly but use several independent pathways via intermediary proteins. Their main mode of anchoring appears to be to the actin cytoskeleton; this can be through interactions with actin-binding proteins such as ~-actinin and spectrin, or more indirectly through scaffold proteins (e.g. via PSD-95-mediated interactions). The involvement of the actin cytoskeleton in NMDA receptor localization is evidenced by the fact that depolymerization of F-actin by latrunculin A causes a 40% reduction in the number of synaptic NMDA receptor clusters, without affecting NMDA receptor clustering (Allison et al., 1998). 195

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The partial effect of actin depolymerization on synaptic clustering of NMDA receptors is consistent with multiple complex interactions between NMDA receptors and the cytoskeleton. NMDA receptors may also associate with microtubules, albeit indirectly via PSD-95 and CRIPT and MAP1A, and they may even interact with neurofilaments via less well-defined mechanisms. The relevance of microtubules and neurofilaments at postsynaptic sites is still controversial. Biochemically, AMPA receptors are easier to solubilize than NMDA receptors, and they appear to be more dynamically regulated in their subcellular targeting, with substantial amounts present in intracellular compartments (Molmir et al., 1993; Baude et al., 1995; Nusser et al., 1998; Petralia et al., 1999). So far, relatively few interactions with cytoskeleton have been uncovered for AMPA receptors or AMPA receptor binding proteins, though the synaptic clustering of AMPA receptors is impaired in latrunculin-A-treated neurons (Allison et al., 1998). Nevertheless, the different subunits of AMPA receptors appear to mediate interactions with distinct sets of cytoplasmic proteins, including the multi-PDZ scaffolds GRIP and ABE The binding of NSF to AMPA receptor GluR2 subunits seems to allude to the dynamic nature of the trafficking and regulation of AMPA receptors. Surprisingly, metabotropic and NMDA-type glutamate receptors may be physically coupled via Shank which bridges Homer to the PSD-95/GKAP complex (Naisbitt et al., 1999; Tu et al., 1999). Such a network of synaptic protein interactions may functionally couple NMDA receptors to the IP3R or ryanodine receptor and thus contribute to the activity-dependent release of Ca 2+ from intracellular stores (Emptage et al., 1999; reviewed in Svoboda and Mainen, 1999). The C-termini of mGluRla/5 bind to the PDZ domain of Shank; thus all postsynaptic GluRs seem to participate in PDZ-based interactions. In addition, by binding to Homer via the PPXXF motif, mGluRs are similar to AMPA and NMDAr subunits in utilizing internal segments of their cytoplasmic tail to associate with non-PDZ proteins. Although each class of glutamate receptor interacts directly with a different set of cytoplasmic proteins, they all seem to act as a membrane node from which emanates a network of specific protein-protein interactions into the cell interior. These networks of proteins may overlap and converge at some points, such as on the Shank family of proteins. The functional consequences of these biochemical interactions need to be determined. In the past several years, the synaptic targeting of glutamate receptors and their associated proteins has been an explosive field in molecular cellular neuroscience, but much remains to be learned. We are at a qualitative phase in the description of the various proteinprotein interactions involving glutamate receptors, and we know little about the functional significance of most of the interacting proteins. A future challenge will be to understand the developmental and activity-dependent regulation of these receptor-associated complexes, and ultimately to discover the roles of these protein interactions in glutamate receptor regulation, synapse development and synaptic plasticity.

7. ACKNOWLEDGEMENTS Supported by the National Institutes of Health (M.W. and M.S.). M.S. is an Assistant Investigator of the Howard Hughes Medical Institute.

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CHAPTER VII

Enzymes responsible for glutamate synthesis and degradation T. KANEKO

1. I N T R O D U C T I O N L-Glutamate and possibly L-aspartate are well known to be major excitatory neurotransmitters in the mammalian central nervous system (CNS). They cause ionotropic glutamate receptors to produce excitatory postsynaptic potentials, and metabotropic glutamate receptors to control intracellular signal transmissions such as calcium/phosphatidyl inositol and cyclic AMP signalings. These excitatory amino acids are, on the other hand, general substrates for living cells which use the amino acids as resources for energy metabolism, raw materials for protein synthesis, and so on. Thus, there could be an overlap between the synthesis and degradation of transmitter glutamate/aspartate and that of metabolic amino acids (cf. Chapters I and II). L-Glutamate is synthesized directly from L-glutamine, 1-pyrroline-5-carboxylate (P5C) or c~-ketoglutarate (2-oxoglutarate) in the CNS (Fig. 1; for review, cf. Shank and Campbell, 1983). The formation of glutamate from glutamine is an energy-saving process catalyzed by phosphate-activated glutaminase (PAG), which is thought to play a major role in the production of transmitter glutamate. P5C is derived from ornithine through glutamic semialdehyde

~

OAT

I

AS

,AAA

sparaginase

P5CDH

/

GDH etc.

PAG/ /GS ~' /

,

GAD

TCA cycle Fig. 1. Metabolic map for synthesis and metabolism of glutamate and aspartate. AAT = aspartate aminotransferase;

AS - asparagine synthetase; GAD -- glutamic acid decarboxylase; GDH -- glutamate dehydrogenase; GS -glutamine synthetase; OAT - ornithine D-aminotransferase; P5CDH = 1-pyrroline-5-carboxylatedehydrogenase; PAG = phosphate-activated glutaminase; PO = proline oxidase; TCA = tricarboxylic acid.

Handbook of Chemical Neuroanatom); Vol. 18." Glutamate O.P. Ottersen and J. Storm-Mathisen, editors (~ 2000 Elsevier Science B.V. All rights reserved.

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by the catalysis of ornithine ~-aminotransferase (OAT) or from proline by proline oxidase (PO), and then converted to glutamate by P5C dehydrogenase (P5CDH). Although OAT immunoreactivity was reported to be located in neurons (Kasahara et al., 1986), P5CDH and PO activities were detected only in astrocytes or Bergmann glial cells (Thompson et al., 1985). et-Ketoglutarate is transformed to glutamate through reductive amination catalyzed by the reverse reaction of glutamate dehydrogenase (GDH). ~-Ketoglutarate is also converted to glutamate through transamination reaction catalyzed by several aminotransferases such as aspartate aminotransferase (AAT) and alanine aminotransferase. Brain alanine aminotransferase activity is much slower than AAT activity (for review, cf. Benuck and Lajtha, 1975), and the activity of synaptosomal alanine aminotransferase is much slower than that of glutaminase (Erecinska et al., 1994), suggesting that alanine aminotransferase plays no major role in neurotransmitter synthesis. In the following sections, I will first review the distribution of glutaminase in the brain, then describe glutamate synthesis and metabolism in glial cells, and finally discuss the role of glutamate and AAT in GABAergic neurons.

2. DISTRIBUTION OF GLUTAMINASE IN THE NERVOUS SYSTEM

It has been established that the transmitter pool of glutamate is preferentially supplied from glutamine (Bradford et al., 1978; Hamberger et al., 1979a,b; Thanki et al., 1983). Bradford et al. (1978) reported that stimulus-released glutamate from cortical synaptosomes was derived principally (80%) from glutamine after co-incubation with [14C]glucose and [3H] glutamine as resources for glutamate. Using hippocampal slices, Hamberger et al. (1979a,b) demonstrated that glutamine was superior to glucose or pyruvate as the precursor of glutamate that was released in a calcium-dependent manner by application of a high concentration of potassium. Thanki et al. (1983) added the in vivo finding that the rat sensorimotor cortex increased the uptake of superfused [~4C]glutamine and the incorporation of radioactivity into the released glutamate during the stimulation period with a high concentration of potassium or tityustoxin. These biochemical findings have indicated that transmitter glutamate is principally formed by hydrolysis of glutamine, and stimulated the immunocytochemical studies to localize the enzyme responsible for glutamine hydrolysis, phosphate-activated glutaminase (L-glutamine amidohydrolase, EC 3.5.1.2), in the nervous system. Glutaminase-like immunoreactivity was first visualized in the auditory nerve neurons (Altschuler et al., 1984) and then in the neocortical neurons (Donoghue et al., 1985) using an antiserum against rat kidney glutaminase. Since then, using the same antiserum, several research groups reported the localization of glutaminase in several brain regions (Altschuler et al., 1985; Cangro et al., 1985; Beitz et al., 1986; Magnusson et al., 1986; Monaghan et al., 1986; Wenthold et al., 1986; Clements et al., 1987; Beitz and Ecklund, 1988; Turman and Chandler, 1994). In 1987, brain glutaminase was purified from rat tissue and monoclonal antibodies were produced against the brain enzyme (Kaneko et al., 1987b, 1988b). The distribution of glutaminase in the mammalian brain was then reported by using the monoclonal antibodies (Kaneko et al., 1987b, 1989; Kaneko and Mizuno, 1988, 1992a) or by applying a polyclonal antibody against brain glutaminase (Akiyama et al., 1990). Both the monoclonal and polyclonal antibodies against brain glutaminase absorbed more than 90% of glutaminase activity of the rat and human brains in a dose-dependent manner (Kaneko et al., 1987b, 1988b; Akiyama et al., 1990). This indicates that these antibodies 204

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are sufficient markers for brain glutaminase activity. Recently, a cDNA encoding glutaminase has been cloned from a rat brain library (Banner et al., 1988; Shapiro et al., 1991), and in situ hybridization histochemistry with oligonucleotide probes has revealed that many neurons including pyramidal cells in the cortex showed signals for glutaminase mRNA (Najlerahim et al., 1990), which supported the immunocytochemical findings. The distribution of glutaminase-like immunoreactivity which was visualized in the rat CNS with the monoclonal antibody against brain glutaminase is summarized in Table 1 and Fig. 2. Fig. 3 shows some examples of glutaminase-immunoreactive neuronal cell bodies in the brain. 2.1. FOREBRAIN REGIONS Glutaminase-immunoreactive neuronal cell bodies were observed in the mitral cell layer, in the outer part of the external plexiform layer, and periglomerular regions of the olfactory bulb (Figs. 2a,b and 3b; Kaneko and Mizuno, 1992b). The neuropil in the external plexiform layer and that in the glomeruli were intensely immunolabeled for glutaminase. In the cerebral neocortex, glutaminase immunoreactivity was located mainly in pyramidal neurons (Figs. 3a, 4b and 7a; Donoghue et al., 1985; Kaneko et al., 1987a, 1995; Kaneko and Mizuno, 1988; Akiyama et al., 1990). Layer V and layer VI pyramidal neurons were stained more intensely than pyramidal neurons in layers II-IV, although some layer VI pyramidal neurons were immunonegative for glutaminase (Kaneko et al., 1995). Glutaminase immunoreactivity has also been observed in cortical neurons of non-pyramidal shape (Akiyama et al., 1990), and in spiny stellate cells located at cortical layer VI (Kaneko and Mizuno, 1996). However, no glutaminase-positive neocortical neurons were immunoreactive for markers of the GABAergic subpopulation, such as parvalbumin, choline acetyltransferase, corticotropin-releasing factor, cholecystokinin, somatostatin, neuropeptide Y and vasoactive intestinal polypeptide (Fig. 4a; Kaneko et al., 1992b). Furthermore, neither GABA- nor glutamic acid decarboxylase (GAD)-immunoreactivity was detected in glutaminase-positive neocortical neurons (Fig. 7c; Kaneko and Mizuno, 1994). These results strongly indicate that glutaminase immunoreactivity is a good marker for glutamatergic, excitatory neurons in the neocortex. In the mesocortical and paleocortical areas and the hippocampal formation, many pyramidal cells and granule cells also showed glutaminase immunoreactivity (Altschuler et al., 1985; Kaneko and Mizuno, 1988). In layer Ia of the piriform cortex, where inputs from the olfactory bulb terminate, neuropil was intensely labeled for glutaminase (Fig. 2d-g). Neuropil of the stratum lacunosum-moleculare of hippocampal CA3 was also intensely immunoreactive for glutaminase (Fig. 2g,h). In the rat neostriatum and accumbens nucleus, glutaminase immunoreactivity was distributed in neuropil with a mosaic organization (Fig. 2d-g; Kaneko and Mizuno, 1992a). Neuropil with strong glutaminase immunoreactivity corresponded to 'patch' regions showing poor immunoreactivity for choline acetyltransferase. Under the electron microscope, glutaminase immunoreactivity was located not only in mitochondria but also in the cytoplasm of axon terminals of the striatum (Aoki et al., 1991). Weak glutaminase immunoreactivity in 'matrix' neuropil was considered to derive from the neocortical projection neurons, whereas intense immunoreactivity in 'patch' regions was intrinsic in origin (Kaneko and Mizuno, 1992a). Actually some medium-size cell bodies with glutaminase immunoreactivity were found in 'patch' regions. Furthermore, all large neostriatal neurons with immunoreactivity for choline acetyltransferase showed glutaminase immunoreactivity. In the basal forebrain, almost all large cholinergic neurons also displayed glutaminase immunoreactivity. These results suggest that the cholinergic neurons in the basal ganglia and basal forebrain regions use glutamate 205

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TABLE 1. Distribution of glutaminase immunoreactivity in the nervous system Glutaminase immunoreactivity

Region

Neuropil

Perikarya

Main olfactory bulb Glomerular layer External plexiform layer Mitral cell layer Internal granular layer Accessory olfactory bulb Anterior olfactory nucleus Olfactory tubercle Islands of Calleja Nucleus of the lateral olfactory tract

++~+++ +§ § § §247 § ++ §

+ + + scattered §247 § scattered §247 .i.-",i,,i,

Neocortex Layer I Layer II-IV Layer V Layer VI

§247 §247 § ++

§ §247 +'~++

Mesocortical areas Layer I Layer II-IV Layer V Layer VI

++ ++ ++ ++

§ + +'~++ +'~++

Piriform cortex Layer I Layer II Layer III

+++ + §

+ § 2 4 7.

Entorhinal cortex Layer I Layer II Layer III Layer IV Layer V Layer VI Tenia tecta Indusium griseum

§247 § §247 § § § ++ ++

§ §247 § §247 § ++ +

Ammon's horn CA1, stratum CA1, stratum CA1, stratum CA1, stratum CA3, stratum CA3, stratum CA3, stratum CA3, stratum CA3, stratum

+ + + § §247 § §247 §247 §247247

+ §247247247247 -

+~++ + + + ++

+ +++ ++ -

oriens pyramidale radiatum lacunosum-moleculare oriens pyramidale lucidum radiatum lacunosum-moleculare

Dentate gyrus Molecular layer Granule cell layer Polymorph layer Subiculum Presubiculum

206

scattered

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Enzymes responsible for glutamate synthesis and degradation TABLE 1 (continued)

Region

Glutaminase immunoreactivity

Neuropil

Perikarya

Parasubiculum Postsubiculum

+ +

+ +

Septal and basal forebrain regions Medial septal nucleus Lateral septal nucleus Septofimbrial nucleus Nucleus of the diagonal band Substantia innominata Bed nucleus of the stria terminalis

+ + ++ + + +

+ + + ++ + +

Basal ganglia Neostriatum Accumbens nucleus Globus pallidus Ventral pallidum Entopeduncular nucleus Subthalamic nucleus Endopiriform nucleus Claustrum

++ ++ + + + ++ + ++

+ scattered + + + +~++ ++ +--~++ +

Amygdala Cortex-amygdala transition zone Anterior amygdaloid area Anterior cortical amygdaloid nucleus Posteromedial cortical amygdaloid nucleus Posterolateral cortical amygdaloid nucleus Medial amygdaloid nucleus Basolateral amygdaloid nucleus Basomedial amygdaloid nucleus Lateral amygdaloid nucleus Central amygdaloid nucleus Intercalated amygdaloid nucleus Amygdalohippocampal area

+ + +~++ + + + ++ + + + + +

+ + +

Habenula Medial habenular nucleus Lateral habenular nucleus

++ +

4++

Thalamus Anterodorsal nucleus Anteroventral nucleus Anteromedial nucleus Mediodorsal nucleus Laterodorsal nucleus Lateroposterior nucleus Ventrolateral nucleus Ventroposterolateral nucleus Ventroposteromedial nucleus Thalamic gustatory nucleus Posterior nuclear group Gelatinosus (or submedius) nucleus Centrolateral nucleus Centromedial nucleus Paracentral nucleus

+++ ++~+++ ++ + ++ ++ + + + + ++ + ++ ++ ++

++ + ++ + ++ ++ ++ ++ ++ ++ ++ + ++ ++ ++

++~+++ + •

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TABLE 1 (continued)

Region

Glutaminase immunoreactivity Neuropil

Perikarya

++ + + ++ +++ + + ++ +++

++ ++ + + + + + ++ ++

+ + + + ++

++ + + ++ + ++ ++

+ + + ++ + ++ + + + scattered ++ + + + scattered + + + + + ++ + +,-~++ ++

Pretectum Nucleus of the optic tract Posterior pretectal nucleus Medial pretectal nucleus Olivary pretectal nucleus Precommissural nucleus Anterior pretectal nucleus

+ + ++ +++ + +

++ + ++ + + +

Terminal nuclei of the accessory optic tract Medial terminal nucleus Lateral terminal nucleus Dorsal terminal nucleus

++ ++ +

Superior colliculus Zonal layer Superficial gray layer Optic nerve layer Intermediate gray layer Intermediate white layer Deep gray layer Deep white layer

++ ++ + + + + +

Parafascicular nucleus Subparafascicular nucleus Reuniens nucleus Paratenial nucleus Paraventricular nucleus Rhomboid nucleus Thalamic reticular nucleus Lateral geniculate nucleus Medial geniculate nucleus Zona incerta Preoptic region and hypothalamus Lateral preoptic area Medial preoptic area Medial preoptic nucleus Magnocellular preoptic nucleus Suprachiasmatic nucleus Supraoptic nucleus Anterior hypothalamic area Lateral hypothalamic area Periventricular nucleus Paraventricular hypothalamic nucleus Arcuate nucleus Dorsal hypothalamic area Ventromedial hypothalamic nucleus Dorsomedial hypothalamic nucleus Compact part Tuber cinereum Posterior hypothalamic area Premammillary nucleus Supramammillary nucleus Medial mammillary nucleus Lateral mammillary nucleus

208

m

+ + ++ + + + +

Enzymes responsible for glutamate synthesis and degradation

Ch. VII

TABLE 1 (continued) Region

Glutaminase immunoreactivity Neuropil

Perikarya

Parabigeminal nucleus Periaqueductal gray (central gray) Edinger-Westphal nucleus Darkschewitsch nucleus Interstitial nucleus of Cajal Oculomotor nucleus Trochlear nucleus Peripeduncular nucleus Ventral tegmental area of Tsai Substantia nigra pars compacta Substantia nigra pars reticulata Substantia nigra pars lateralis Red nucleus maganocellular part parvocellular part Retrorubral field Inferior colliculus Nucleus of the brachium of the inferior colliculus Subbrachial nucleus Cuneiform nucleus Mesencephalic trigeminal nucleus

++ + + ++ ++ ++ ++ + + ++ + +

++ + ++ ++ ++ + some small cells + ++ +~++ +,~++ +--~++ ++

+ + + + + + ++ +

+~++ + scattered + + + + + ++

Rostral linear raphe nucleus Caudal linear raphe nucleus Median raphe nucleus Dorsal raphe nucleus

+ + + +

+ +~++ +§ ++

Mesencephalic reticular formation Pedunculopontine tegmental nucleus Microcellular tegmental nucleus Interpeduncular nucleus

+ + +~++ +++

Lateral parabrachial nucleus Medial parabrachial nucleus Locus coeruleus Supratrigeminal region Peritrigeminal region Nucleus of the lateral lemniscus Laterodorsal tegmental nucleus Posterodorsal tegmental nucleus Pontine raphe nucleus Dorsal tegmental nucleus Ventral tegmental nucleus Superior olivary complex Nucleus of the trapezoid body Trigeminal motor nucleus Principal sensory trigeminal nucleus Dorsal portion Pontine nuclei Pontine tegmental reticular nucleus of Bechterew Nucleus k Pontine reticular formation

++ ++ ++ + + ++ + + ++ ++ + ++ + + + + +++ +++ +++ +

+ + + + + ++ + + + + ++ ++ ++ + + + + scattered +++ + + + dense + + + dense +++ +

Cerebellar cortex Molecular layer

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TABLE 1 (continued)

Region

Glutaminase immunoreactivity Neuropil

Purkinje cell layer Granule cell layer Deep cerebellar nuclei

+++

++

Spinal trigeminal nuclei Oral subnucleus Oral subnucleus, dorsomedial part Interpolar subnucleus Caudal subnucleus, lamina I Caudal subnucleus, lamina II Paratrigeminal nucleus Nucleus of the solitary tract Area postrema Abducens nucleus Facial nucleus Paracochlear glial substance Dorsal cochlear nucleus Granule cell layer Ventral cochlear nucleus Posterior ventral cochlear nucleus

+ ++ ++ + + ++ ++ +++ + ++

Vestibular nucleus Superior vestibular nucleus Medial vestibular nucleus Medial vestibular nucleus, ventral part Lateral vestibular nucleus Inferior vestibular nucleus Cell group f Hypoglossal prepositus nucleus Intercalated nucleus Nucleus of Roller Linear nucleus Cell group x Cell group y Nucleus z Paracochlear glial substance Cell group e Supragenual nucleus Lacrimo-nasopalatine nucleus Dorsal motor nucleus of the vagus nerve Ambiguus nucleus Hypoglossal nucleus External cuneate nucleus Cuneate nucleus Gracile nucleus Inferior olivary nuclei Lateral reticular nucleus of the medulla Medullary reticular formation Gigantocellular reticular formation Paramedian reticular formation Raphe magnus nucleus Raphe pallidus nucleus Raphe obscurus nucleus

+++ +++ +++ + ++ +++ +++ +++ +++ +++ +++ +++ + +++ ++ ++ ++ ++ + + +++ ++ + ++~+++ +++ + + + + + +

210

Perikarya

n.e.

+

+++ + + + scattered + scattered +++ + + + +++ + m

++ +++ +++ +++ + + +++ +++ +++ +++ +++ +++ +++ + +++ +++ + + + scattered ++ ++ ++ 4+++ + + +++ + + + + scattered + + + scattered + + +

Enzymes responsible for glutamate synthesis and degradation

Ch. VII

TABLE 1 (continued) Region

Glutaminase immunoreactivity Neuropil

Perikary a

Spinal cord Lamina I Lamina II Lamina III Lamina IV Lamina V Lamina VI Lamina VIII Lamina IX General motoneurons Pudendal motoneurons Lamina X Intermediolateral nucleus Sacral parasympathetic nucleus

+ ++ ++ + + ++ + + n.e. n.e. ++ ++ ++

+ scattered + scattered + scattered + scattered + scattered + +~++ ++ +-~ + + +--~++

Trigeminal g0nglion Vestibular ganglion of Scarpa Spiral ganglion Nodose ganglion Dorsal root ganglia Superior cervical ganglion Celiac ganglion Pelvic ganglion

-

++ ++ ++ ++ ++ -

Intensity of glutaminase immunoreactivity was evaluated using internal standards as in Kaneko et al. (1989): medial geniculate nucleus for intense ( + + + ) neuropil immunoreactivity; paracochlear glial substance for intense ( + + + ) perikaryal immunoreactivity; superficial gray layer of the superior colliculus for moderate (++) neuropil immunoreactivity; Edinger-Westphal nucleus or mesencephalic trigeminal nucleus for moderate (++) perikaryal immunoreactivity; central gray of the midbrain for low (+) neuropil immunoreactivity; parvocellular part of the red nucleus for low (+) perikaryal immunoreactivity; - , negative; n.e., not evaluated.

as well as acetylcholine for their transmitters. M o s t neurons in the subthalamic nucleus showed g l u t a m i n a s e i m m u n o r e a c t i v i t y (Fig. 2h; K a n e k o and Mizuno, 1992a), which result is consistent with the excitatory nature of subthalamic nucleus neurons. In the (external s e g m e n t of the) globus pallidus, e n t o p e d u n c u l a r nucleus (i.e. internal s e g m e n t of the globus pallidus) and substantia nigra pars reticulata, w e a k l y g l u t a m i n a s e - i m m u n o r e a c t i v e neurons were found ( K a n e k o et al., 1990; K a n e k o and Mizuno, 1992a). This suggests either that g l u t a m i n a s e is used to produce G A B A precursor g l u t a m a t e in G A B A e r g i c neurons, or that these nuclei contained a subpopulation of n o n - G A B A e r g i c neurons, such as cholinergic neurons in the globus pallidus, in addition to G A B A e r g i c projection neurons. In the a m y g d a l o i d nucleus, m o d e r a t e l y to intensely g l u t a m i n a s e - i m m u n o r e a c t i v e neurons were found in the basolateral nucleus, and w e a k l y i m m u n o r e a c t i v e neurons were o b s e r v e d in the anterior, anterior cortical and lateral nuclei. 2.2. D I E N C E P H A L I C R E G I O N S In the diencephalic structure, almost all the neurons of the dorsal thalamic nuclei contained g l u t a m i n a s e immunoreactivity, although the intensity of i m m u n o r e a c t i v i t y was different from nucleus to nucleus (Figs. 2 f - i and 3c; K a n e k o and Mizuno, 1988). In layer IV of the rat 211

Ch. VII

T. Kaneko

Fig. 2. Distribution of glutaminase-like immunoreactivity in the central nervous system. The rat brains were fixed and immunostained as reported before (Kaneko et al., 1989). Briefly, rats were fixed at room temperature by transcardial perfusion of 0.2% formaldehyde, 75%-saturated picric acid and 0.1 M sodium-phosphate, pH 7.0, and the brain blocks were further placed for 3 days at 4~ in 2% formaldehyde, 75%-saturated picric acid and 0.1 M sodium-phosphate, pH 7.0. The brain sections (30 Ixm thick) were incubated with 10 Ixg/ml monoclonal anti-glutaminase IgM, MAb-120, then with 10 [xg/ml biotinylated goat anti-glutaminase antibody (Vector), and finally with avidin-biotinylated peroxidase complex (ABC; Vector), and the bound peroxidase was developed brown by reaction for 10-30 min with 0.02% diaminobenzidine-4HC1 (DAB), 0.003% H202 and 50 mM Tris-HC1, pH 7.6. (o) Upper cervical cord. Abbreviations: A1 = primary auditory area; ac = anterior commissure; Acc = accumbens nucleus; AON = anterior olfactory nucleus; BF = barrel field; BLA = basolateral nucleus of the amygdala; CA1 = cornu ammonis 1; CA3 = cornu ammonis 3; cc = corpus callosum; Cg = cingulate area; CPu = caudate-putamen; DCb = deep cerebellar nuclei; DCo = dorsal cochlear nucleus; DG = dentate gyrus; DMV = dorsal motor nucleus of the vagus nerve; ECu = external cuneate nucleus; EP = external plexiform layer; ER = entorhinal cortex; f = fornix; Fa = facial nucleus; fa = facial nerve; fr = fasciculus retroflexus; G1 = glomerular layer; GPe = (external segment of the)

212

Enzymes responsible for glutamate synthesis and degradation

Ch. VII

t

f

ot

0

g

2ram II

III

Fig. 2 (continued). globus pallidus; GPi = e n t o p e d u n c u l a r nucleus (internal s e g m e n t of the globus pallidus); G r -- g r a n u l a r layer; IC = inferior colliculus; icp = inferior cerebellar peduncle; IG -- internal g r a n u l a r layer; IO -- inferior olivary c o m p l e x ; Ip -- i n t e r p e d u n c u l a r nucleus; L D = laterodorsal t h a l a m i c nucleus; L G --- lateral geniculate nucleus; Li -- linear nucleus; 11 = lateral lemniscus; lot -- lateral olfactory tract; L P --- lateroposterior t h a l a m i c nucleus; L R -- lateral reticular nucleus of the m e d u l l a oblongata; M1 = p r i m a r y m o t o r area; m c p -- m i d d l e cerebellar p e d u n c l e ; M D = m e d i o d o r s a l t h a l a m i c nucleus; M G -- m e d i a l geniculate nucleus; M m --- m a m m i l l a r y nucleus; M o -- m o l e c u l a r layer; mt -- m a m m i l l o t h a l a m i c tract; MVe = m e d i a l vestibular nucleus; oc - optic chiasm; OT = olfactory t u b e r c u l u m ; ot -- optic tract; P -- pontine nuclei; Pir -- p i r i f o r m cortex; P T R = p o n t i n e t e g m e n t a l reticular nucleus of B e c h t e r e w ; Re -- red nucleus; RS -- retrosplenial area; S -- septal

213

Ch. VII

T. Kaneko

Mo

9

D

i .

,~

~".~,'i~"~',

P

'."~b~"'.~" ,,.~.r ' ~ :'~ ~,

,

-~,'

:

SO

II'

113

2ram 9

IIII

IIIIIII

I

O LR Fig. 2 (continued). nuclei; S1 = primary somatosensory nucleus; SC = superior colliculus; SCh = suprachiasmatic nucleus; sm = stria medullaris; SN = substantia nigra; SO = superior olivary complex; SOp = supraoptic nucleus; Sol = solitary tract nucleus; SpVe = spinal vestibular nucleus; sr = sensory root of the trigeminal nerve; STh = subthalamic nucleus; SVe = superior vestibular nucleus; V 1 = primary visual area; V2 = secondary visual area; VCo = ventral cochlear nucleus; Vc = caudal subnucleus of the spinal trigeminal nucleus; Vi = interpolar subnucleus of the spinal trigeminal nucleus; Vp = principal nucleus of the trigeminal nerve; XII = hypoglossal nucleus.

214

Enzymes responsible for glutamate synthesis and degradation

Ch. VII

primary somatosensory cortex, intensely glutaminase-immunoreactive barrels were seen (BF in Fig. 2e-g; Kaneko and Mizuno, 1988), indicating that axon terminals of thalamocortical projection neurons were labeled with glutaminase immunoreactivity. Moderate glutaminase immunoreactivity was seen in GABAergic neurons of the thalamic reticular nucleus. Glutaminase may serve as an enzyme supplying GABA-precursor glutamate in the reticular nucleus neurons.

In the preoptic and hypothalamic regions, moderately glutaminase-immunoreactive neurons were found in the magnocellular preoptic, supraoptic, premammillary, paraventricular and mammillary nuclei and dorsal hypothalamic area (Fig. 2f-h). Many weakly immunoreactive neurons were scattered throughout the preoptic and hypothalamic regions. Moderate glutaminase immunoreactivity was observed in neuropil of the parastrial, suprachiasmatic, premammillary and mammillary nuclei and the compact part of the dorsomedial hypothalamic nucleus, although most neuropil in preoptic and hypothalamic regions showed a weak glutaminase immunoreactivity. 2.3. BRAINSTEM AND CEREBELLAR REGIONS In the lower brainstem, the most conspicuous finding was that almost all the precerebellar nuclei sending mossy fibers to the cerebellar cortex contained intensely glutaminase-immunoreactive neurons (Kaneko et al., 1987b, 1989); most neurons in the pontine nuclei (Figs. 2j and 3c; Beitz et al., 1986), the pontine tegmental reticular nucleus of Bechterew (Fig. 2j), the lateral reticular nucleus of the medulla oblongata (Fig. 2n), and the external cuneate nucleus showed intense glutaminase immunoreactivity. In nucleus k, cell groups e, f, x and y, interpolar subnucleus of the spinal trigeminal nucleus, hypoglossal prepositus nucleus, intercalated nucleus, nucleus of Roller, linear nucleus (Fig. 2m), superior and medial vestibular nuclei (Fig. 2k,1), and gigantocellular and paramedian reticular formation, intensely immunoreactive neuronal cell bodies were scattered. The latter nuclei are also known to contain neurons sending mossy fibers to the cerebellar cortex. In the cerebellar cortex, mossy fiber endings in the granular layer were intensely labeled with glutaminase immunoreactivity (Fig. 2k; Wenthold et al., 1986; Kaneko et al., 1987b, 1989; Laake et al., 1999). These results suggest that most mossy fiber inputs to the cerebellum are glutamatergic. In contrast, cerebellar granule cells and neuropil of the molecular layer, i.e. the region where parallel fibers run, showed little or faint immunoreactivity for glutaminase (Wenthold et al., 1986; Kaneko et al., 1987b, 1989), although an mRNA signal for glutaminase was detected on the granule cells (Najlerahim et al., 1990). Three to five days after intraventricular injection of an irreversible inhibitor of glutaminase, 6-diazo-5oxo-L-norleucine, glutaminase immunoreactivity was much enhanced in many brain regions because of the compensatory production of new enzyme proteins (Kaneko et al., 1992a). After this treatment, neuropil in the molecular layer of the cerebellar cortex showed an increase in granular immunoreactivity for glutaminase, suggesting that granule cell-parallel fiber systems use glutaminase to synthesize the transmitter glutamate. Neurons in the deep cerebellar nuclei were weakly to moderately immunoreactive for glutaminase (Fig. 21; Monaghan et al., 1986; Kaneko et al., 1989). Since neurons in the inferior olivary complex were embedded in moderately immunoreactive neuropil, it was difficult to identify glutaminase-immunoreactive neurons in the complex (Fig. 2m,n; Kaneko et al., 1987b, 1989). However, at a developmental stage, postnatal days 0 to 21, clearly immunoreactive neuronal cell bodies were recognized in the inferior olivary complex (Kaneko and Mizuno, 1992b). Weak glutaminase immunoreactivity was observed in neurons of the red 215

Ch. VII

T. Kaneko

~4

Fig. 3. Glutaminase-immunoreactive neurons in layer V of the neocortex (a), mitral cell layer of the olfactory bulb (b), lateral geniculate nucleus (c) and pontine nuclei (d). The method for immunostaining is described in the legend of Fig. 1. EP = external plexiform layer; IG = internal granular layer; M -- mitral cell layer. Modified from Kaneko (1991).

nucleus, including rubrospinal projection neurons (Fig. 2i; Beitz and Ecklund, 1988; Kaneko et al., 1989). Intense to moderate glutaminase immunoreactivity was observed in neuropil of the medial and olivary pretectal nucleus, superficial layer of the superior colliculus, and medial and lateral terminal nuclei of the accessory optic tract, suggesting that retinal inputs to these nuclei use glutamate as their transmitters as well as retinogeniculate inputs. Moderate immunoreactivity was also found in neuropil of the oculomotor nucleus, trochlear nucleus, Darkschewitsch nucleus and interstitial nucleus of Cajal. Neuropil of the solitary tract nucleus and that of the lateral and parabrachial nuclei showed moderate immunoreactivity for glutaminase. This result, together with the presence of glutaminase-positive neurons in the nodose ganglion (Li et al., 1996), suggests that glutaminase and glutamate is associated with information 216

Enzymes responsible for glutamate synthesis and degradation

Ch. VII k ~

'

L~

,

g

Fig. 4. Glutaminase-immunoreactive neurons in the rat (a) and human cerebral cortex (b). (a) Glutaminaseimmunoreactive neurons and parvalbumin-immunoreactive ones were stained green and red, respectively, by the double immunofluorescence method as described before (Kaneko et al., 1992b). Rat sections were incubated with anti-glutaminase mouse IgM and anti-parvalbumin mouse IgG, then with biotinylated anti-mouse IgG Fc portion donkey antibody, and finally with Texas Red-conjugated avidin D and fluorescein-labeled anti-mouse IgM donkey antibody. (b) Many pyramidal cells including Betz cells (arrow) show glutaminase immunoreactivity in the human motor cortex (Akiyama et al., 1990). The human cortical sections were incubated with a rabbit serum against rat brain glutaminase, with biotinylated anti-rabbit IgG goat antibody, and with ABC. The bound peroxidase was developed blue black by reaction with 0.1% DAB, 1% nickel ammonium sulfate, 0.05% imidazole and 0.00016% H202 in 50 mM Tris-HC1, pH 7.6.

transmission of taste and visceral senses. In contrast, many neuronal cell bodies in the central part of the dorsal column nuclei, dorsal horn of the spinal cord, principal trigeminal nucleus and caudal subnucleus of the spinal trigeminal nucleus showed no or faint glutaminase immunoreactivity, raising the question of what is used for their excitatory transmission. Monoaminergic neurons, such as dopaminergic ones in the substantia nigra pars compacta, noradrenergic ones in the locus ceruleus and serotonergic ones in the raphe nuclei, showed weak to moderate immunoreactivity for glutaminase, although no dopaminergic neurons in the olfactory bulb displayed glutaminase immunoreactivity (Kaneko et al., 1990; Minson et al., 1991). These results suggest that glutamate is used as a transmitter or a precursor of GABA by brainstem monoaminergic neurons, but not by monoaminergic ones in the olfactory bulb. 2.4. SPINAL CORD AND PERIPHERAL NERVES In the spinal cord, neuropil of laminae II, III, VI and X were moderately immunoreactive for glutaminase (Fig. 2o). Weakly immunoreactive neurons were found in laminae I, IV-IX, 217

Ch. VII

T. Kaneko

and moderately immunoreactive neurons were scattered in lamina X. Neuropil and neuronal cell bodies of the intermediolateral sympathetic and sacral parasympathetic nuclei showed moderate immunoreactivity (Senba et al., 1991; Chiba and Kaneko, 1993). In the cranial parasympathetic nuclei, the dorsal motor nucleus of the vagus nerve and lacrimo-nasopalatine nucleus also contained moderately immunoreactive neuronal cell bodies and neuropil (Kaneko et al., 1989; Senba et al., 1991). Thus, many autonomic preganglionic neurons appear to be regulated by glutamatergic inputs and use glutamate by themselves. Glutaminase immunoreactivity was also found in some neurons emitting ascending fibers such as spinomesencephalic tract cells (Yazierski et al., 1993). In the dorsal root, trigeminal and nodose ganglia (Cangro et al., 1985; Miller et al., 1993; Li et al., 1996), in the vestibular ganglion of Scarpa (Kaneko et al., 1989), and in the spiral ganglion of the cochlea (Altschuler et al., 1984), moderately glutaminase-immunoreactive neurons were found. These findings indicate that glutaminase is generally used for transmitter synthesis by many kinds of sensory afferent neurons. Furthermore, the size of the immunoreactive cell bodies was widely distributed from small to large in the dorsal root and trigeminal ganglia, suggesting that somatosensory input neurons of various modes, such as pain, touch sense and joint sense, are glutamatergic. In contrast, autonomic ganglion neurons such as the superior cervical, pelvic and celiac ganglia did not show immunoreactivity for glutaminase (Li et al., 1996). 2.5. RETINA Takatsuna et al. (1994) reported the distribution of glutaminase immunoreactivity in the guinea pig, rat and mouse retinae. Many ganglion, bipolar and amacrine cells and possibly horizontal cells showed moderate glutaminase immunoreactivity. In addition, almost all bipolar cells containing protein kinase C were immunoreactive for glutaminase, suggesting that the majority of glutaminase-immunoreactive bipolar cells were of the ON type (Griinert and Martin, 1991). Intense glutaminase immunoreactivity was observed in neuropil of the inner and outer plexiform layers and around the outer limiting membrane. Weak to moderate immunoreactivity was seen in the outer nuclear layer and inner and outer segments of photoreceptors. 2.6. NON-NEURAL DISTRIBUTION OF GLUTAMINASE Although glutaminase is mainly localized in neuronal cells and processes and is not found in small glial cells in most brain regions such as the corpus callosum, Aoki et al. (1991) detected glutaminase immunoreactivity in glial, probably astrocytic processes of the striatal tissue by using electron microscopy. Takatsuna et al. (1994) also observed glutaminase immunoreactivity in processes of Miiller cells (retinal astrocyte-like cells) of the guinea pig retina. These findings suggest an extraneuronal, supposedly metabolic role of glutaminase in glial cells of the brain and retina. For instance, in the kidney, which contains high glutaminase activity, glutaminase is known to play a role in release of ammonia from blood-derived glutamine into urine (for review, cf. Curthoys and Watford, 1995). However, astrocytes and Miiller cells are well known to synthesize glutamine from glutamate and ammonia through an ATP-consuming process catalyzed by glutamine synthetase as described below. Thus, if astrocytes show an activity to degrade glutamine to glutamate and ammonia, it raises the question as to how those couteracting glutaminase and glutamine synthetase activities are decoupled in astrocytes. Otherwise astrocytes might consume all their energy in a reverberating metabolic cycle between glutamine and glutamate/ammonia without producing 218

Enzymes responsible for glutamate synthesis and degradation

Ch. VII

any useful substances. Further studies are necessary to solve this question, although there might be a sort of metabolic sequestration between glutaminase and glutamine synthetase activities in astrocytes.

3. GLUTAMATE SYNTHESIS AND METABOLISM IN GLIAL CELLS Almost at the same time when glutamine was proven to be a preferential precursor for transmitter glutamate, a synthetic enzyme for glutamine, glutamine synthetase (GS; L-glutamate:ammonia ligase [ADP-forming], EC 6.3.1.2), was revealed to be expressed in astrocytes (Martinez-Hernandez et al., 1977; Norenberg, 1979; Norenberg and MartinezHernandez, 1979). Although it has recently been reported that GS immunoreactivity is not only located in astrocytes but also in oligodendrocytes (D'Amelio et al., 1990; Tansey et al., 1991; Miyake and Kitamura, 1992), astrocytes are considered to be main production sites of glutamine, and form the 'glutamine cycle' coupled with neurons employing glutaminase to produce glutamate (Hamberger et al., 1979a,b; Hertz, 1979; Shank and Aprison, 1981). In the 'glutamine cycle', astrocytes take up glutamate released from neurons, convert it to glutamine at the cost of energy, and then supply glutamine to extracellular fluid to maintain a high concentration (about 0.3 raM) of glutamine in the extracellular fluid (Fig. 5). On the other hand, neurons consume glutamine to produce transmitter by an energy-free process catalyzed by glutaminase. Thus, excitatory neurons, so to speak, can use free glutamine as they like. This scheme has largely been supported not only by the recent immunocytochemical studies for GS and glutaminase, but also by those for glutamate and glutamine (see Chapter 1). Recently, immunocytochemical and in situ hybridization histochemical studies revealed that some plasma membrane glutamate transporters, GLT1 and GLAST, were expressed in astrocytes (Danbolt et al., 1992; Levy et al., 1993; Rothstein et al., 1994; Lehre et al., 1995; Schmitt et al., 1997), supporting the 'glutamine

Excitatory Nerve Ending

Capillary Astrocyte (Small Glutamate Pool)

(-

Receptor

h

Fig. 5. Glutamine cycle formed between excitatory nerve endings and astrocytes and de novo synthesis of glutamate in astrocytes (cf. Chapter I). Modified from Kaneko et al. (1988a). 219

Ch. VII

T. Kaneko

cycle' hypothesis by showing astrocytes armed with an uptake mechanism of the released glutamate. A long time has passed since compartmentation of tricarboxylic acid (TCA) cycle-glutamate/glutamine metabolism was proposed in the mammalian CNS (for review, cf. Berl et al., 1975). In ammonia metabolism associated with glutamate/glutamine metabolism, the presence of at least two compartments has been postulated on the basis of labeling patterns after intracarotid administration of isotope (15N)- or radioisotope (13N)-labeled ammonia. Berl et al. (1962) and Cooper et al. (1979) found that the specific activity incorporated into the 0~-amino group of glutamine was several times higher than that of glutamate. To explain this result, they postulated the presence of at least two small and large glutamate compartments in the brain; blood-borne ammonia preferentially entered a small glutamate compartment where glutamate was rapidly metabolized to glutamine, whereas the ammonia was not utilized by a large, metabolically inactive compartment. Since Cooper et al. (1979) observed the incorporation of radioisotope-labeled ammonia into the 0~-amino group of glutamine at the normal level of blood ammonia, it is likely that a part of glutamate in the small compartment is de novo synthesized via reductive amination of 0t-ketoglutarate in a physiological condition. Thus, 0t-ketoglutarate reductive amination activity is considered to be a key enzyme of the small glutamate compartment. The reverse reaction of glutamate dehydrogenase (GDH; L-glutamate-NAD(P) + oxidoreductase [deaminating], EC 1.4.1.3) is 0t-ketoglutarate reductive amination activity. Since the antibody to GDH absorbed more than 95% of c~-ketoglutarate reductive amination activity in the brain (Kaneko et al., 1988a), the small glutamate compartment is characterized by the

Fig. 6. Cellular colocalization of glutamine synthetase (GS) and glutamate dehydrogenase (GDH) in the neostriatum. Each number indicates the same glial cell showing both GDH and GS immunoreactivities. The figure is modified from Kaneko et al. (1988a), where the method for double staining is described. Briefly, GS was immunolabeled with anti-GS rabbit serum and fluorescein-labeled anti-rabbit IgG antibody, and after blocking the sections with normal rabbit serum GDH was visualized by the immunoperoxidase method with biotinylated anti-GDH rabbit IgG and ABC.

220

Enzymes responsible for glutamate synthesis and degradation

Ch. VII

presence of GDH. Immunoreactivity for GDH has been reported to be mainly localized in astrocytes (Aoki et al., 1987a,b; Kaneko et al., 1987a, 1988a; Wenthold et al., 1987; Madl et al., 1988; Rothe et al., 1990, 1994), although a recent in situ hybridization histochemical work showed the neuronal or oligodendrocytic presence of GDH mRNA in addition to a strong astrocytic presence (Schmitt and Kugler, 1999). Since GS and GDH is colocalized in astrocytes (Fig. 6; Kaneko et al., 1988a) to enzymically couple with each other, glutamate which is newly synthesized from 0~-ketoglutarate and ammonia may be preferentially converted to glutamine. These results indicate that astrocytes are the substance of the small glutamate compartment, and that they de novo synthesize glutamate to supply it to the 'glutamine cycle' (Fig. 5).

4. GLUTAMATE AND AAT IN GABA SYNTHESIS Glutamate works not only as an excitatory neurotransmitter but also as the immediate precursor of inhibitory transmitter GABA. Since 0~-ketoglutarate reductive amination activity, which is catalyzed by GDH, and P5C dehydrogenase activity have been immunocytochemically or histochemically shown to be localized in astrocytes (Figs. 6 and 7d; Thompson et al., 1985), glutaminase activity or 0~-ketoglutarate aminotransferase activity, of glutamate-synthesizing enzymic activities indicated in Fig. 1, is considered as a candidate for a direct supplier of GABA-precursor glutamate in the CNS. It has been reported that GABA is formed from 0~-ketoglutarate in synaptosomes of rat brain (Shank and Campbell, 1984a,b), and from glutamine in the cerebral cortex (Tapia and

.._..--__.---, i,

,

~,

ii,

,;~-~ , ~

!t 9

;

,,

- . 2 o :,-:---: ...." . . "

Fig. 7. Glutaminase, mitochondrial and soluble aspartate aminotransferases and glutamate dehydrogenase immunoreactivities in the cerebral neocortex of the rat. The figure is modified from Kaneko and Mizuno (1994). PAG = phosphate-activated glutaminase.

221

b~

TABLE 2. sAAT immunoreactivity in the mammalian retina Species

Rat a Rat b Rat c Rat d Guinea pig e colchicine-treated Guinea pig f Cat g Monkey f Human b

Outer segments layer

Inner segments layer _

-

a Recasens and Delaunoy, 1981. b Brandon and Lam, 1983. c Lin et al., 1983. d Inagaki et al., 1985. e Altschuler et al., 1982. f Mosinger and Altschuler, 1985. g Bolz et al., 1985. h Not described clearly.

Outer nuclear layer m

§ cone § + + cone § 2 4 7cone §247

§ cell body § cell body _9 § cell body

§

-

+ cone

§ cell body

-

Outer plexiform layer _

§ endfoot § § endfoot +§ §247 +§ +-t-

Inner nuclear layer + + cell body + bipolar amacrine + cell body § bipolar amacrine horizontal § cell body § cell body § amacrine § cell body § 2 4 7bipolar amacrine § 2 4 7bipolar amacrine

Inner plexiform layer

§247 + §247 §247 §247 §247

Ganglion cell layer

Mfiller cells

-

§

__

__9 h

--9

--9

§ 2 4 7cell body § cell body

_9 _9

9

--9

§ -

§247 §247

§

§ 2 4 7

Enzymes responsible for glutamate synthesis and degradation

Ch. VII

Gonzfilez, 1978; Bradford et al., 1983; Thanki et al., 1983; Ward et al., 1983) and hippocampus (Reubi, 1980; Bradford et al., 1983; Ward et al., 1983; Szerb, 1984). Thus, glutamate which is formed by transamination of ~-ketoglutarate or by hydrolysis of glutamine is considered to be an immediate precursor of GABA. Glutaminase is, however, localized principally in excitatory neurons, although glutaminase is found in a limited number of GABAergic neurons such as thalamic reticular nucleus neurons in the CNS (Kaneko and Mizuno, 1988). In particular, in the cerebral neocortex, glutaminase immunoreactivity has been proven to be absent in GABAergic interneurons (Kaneko et al., 1992b; Kaneko and Mizuno, 1994). Thus, in most GABAergic cortical neurons, it is unlikely that glutamate formed from glutamine is the immediate precursor of transmitter GABA. It is probable that, in the biochemical works, glutamate derived from glutamine was used indirectly as a remote precursor of GABA through a metabolic link formed with astrocytes as shown in Fig. 10. It is known that the highest transamination activity with ~-ketoglutarate in the brain is catalyzed by aspartate aminotransferases (AAT; L-aspartate:2-oxoglutarate aminotransferase, EC 2.6.1.1; Benuck et al., 1971, 1972; Johnson, 1972). AAT activity in the brain is separated into soluble (cytosolic) and mitochondrial isoenzymes, sAAT and mAAT (for review, cf. Benuck and Lajtha, 1975). Although AAT activity has been detected by classical histochemistry (for review, cf. Lewis and Stoward, 1991; for a recent study of it in the brain, cf. Kugler, 1987), immunocytochemistry with anti-sAAT and anti-mAAT antibodies have made it possible to differentially localize sAAT and mAAT in the nervous system. sAAT immunoreactivity has extensively been studied in the mammalian retina (Recasens and Delaunoy, 1981; Altschuler et al., 1982; Brandon and Lam, 1983; Lin et al., 1983; Bolz et al., 1985; Inagaki et al., 1985; Mosinger and Altschuler, 1985), but the results were somewhat inconsistent with one another (Table 2). Although this inconsistency appears due partly to species differences, there are still rather large differences in the same species. In the rat retina, three reports said that no sAAT immunoreactivity was found in ganglion cells, but another described many clearly immunoreactive neurons in ganglion cell layers, sAAT immunoreactivity was also detected in axon terminals of the auditory nerve (Altschuler et al., 1981) and in dorsal root ganglion neurons (Inagaki et al., 1987). Although sAAT appeared to be used by excitatory neurons in the peripheral nervous system, sAAT-immunoreactive central neurons were intrinsic, probably GABAergic, neurons such as periglomerular and granule cells of the olfactory bulb (Recasens and Delaunoy, 1981; Kamisaki et al., 1984), stellate and basket cells of the cerebellum (Kamisaki et al., 1984; Wenthold et al., 1986) and non-pyramidal neurons in the hippocampus (Altschuler et al., 1985) and cerebral cortex (Figs. 7c and 8a,a'; Donoghue et al., 1985; Kaneko and Mizuno, 1994). In the cerebral cortex, 95% of GABA-immunopositive neurons showed sAAT immunoreactivity, but no glutaminase-positive neurons displayed sAAT immunoreactivity (Fig. 8; Kaneko and Mizuno, 1994), indicating that sAAT is specifically expressed in GABAergic neurons. In addition, many glutamic acid decarboxylase (GAD)-immunoreactive axon terminals contained sAAT immunoreactivity, suggesting that sAAT is enzymatically coupled with GAD in the terminals (Fig. 9; Kaneko and Mizuno, 1994). Thus, it is likely, in cortical GABAergic interneurons, that 0~-ketoglutarate is the precursor of GABA-precursor glutamate, and that sAAT catalyzes the reaction synthesizing GABA-precursor glutamate from ~-ketoglutarate (Fig. 10). mAAT was, in contrast to sAAT, found in both glutaminase-immunoreactive and GABAimmunoreactive neurons in the cerebral cortex (Figs. 7b and 8b,b'; Kaneko and Mizuno, 1994). mAAT immunoreactivity has also been found in mitral, tufted and granule cells and glomeruli of the olfactory bulb (Recasens and Delaunoy, 1981; Kamisaki et al., 1984), Purk223

Ch. VII

T. Kaneko

Fig. 8. Aspartate aminotransferases and glutaminase (PAG) in neocortical GABAergic interneurons. The figure is modified from Kaneko and Mizuno (1994). All sAAT- and some mAAT-immunoreactive neurons are immunoreactive for GABA, but no glutaminase-immunoreactive ones show GABA immunoreactivity. Arrowheads indicate GABA-positive neurons, sAAT, mAAT and glutaminase were stained with primary rabbit or mouse antibodies, biotinylated secondary antibodies and Texas Red-conjugated avidin D, whereas GABA was visualized with antiGABA guinea pig antibody and fluorescein-labeled anti-guinea pig IgG antibody. The photographs in each row were taken at the same site under different excitations for fluorescence. (c') Photograph taken by double exposure.

224

Enzymes responsible for glutamate synthesis and degradation

Ch. VII

Fig. 9. Aspartate aminotransferase immunoreactivity in glutamic acid decarboxylase (GAD)-immunoreactive neuronal processes in the cerebral cortex, sAAT but not mAAT is colocalized with GAD in fine, probably axonal processes (arrows). Rat sections were double-immunostained by incubation with a mixture of anti-sAAT or mAAT rabbit serum and anti-GAD sheep serum, then with biotinylated anti-rabbit IgG donkey antibody, and finally with Texas Red-conjugated avidin and fluorescein-labeled anti-sheep IgG donkey antibody. The photographs in each row were taken at the same site under different excitations. Asterisks in (a) and (a') indicate the unlabeled cell body of a pyramidal neuron.

inje and deep Golgi cells of the cerebellar cortex (Kamisaki et al., 1984), and inner segments, outer plexiform layer and ganglion cells of the retina (Table 3). mAAT is well known to work as a key enzyme with malate dehydrogenase in the malate-aspartate shuttle carrying NADH from cytoplasm to mitochondria for the mitochondrial electron-transport system. Since mAAT is located in virtually all neurons in the cerebral cortex (Figs. 7b and 8b,b'; Kaneko and Mizuno, 1994), mAAT appeared to be associated with the general energy metabolism rather than the specific transmitter-related function in the cortex.

5. C O N C L U D I N G R E M A R K S

It is difficult to specify the functional role of synthetic enzymes for glutamate in the CNS, partly because glutamate is not only an excitatory neurotransmitter but also a general metabolic substrate. However, excitatory neurons appear to use glutamine as a precursor of transmitter glutamate, probably because glutaminase reaction that produces glutamate from 225

Ch. VII

T. Kaneko

TABLE 3. mAAT immunoreactivity in the mammalian retina Species

Outer segments

Inner segments

Outer nuclear layer

Outer plexiform layer

Inner nuclear layer

Inner plexiform layer

Ganglion cell layer

Mtiller cells

Rat a Rat b

_ -

_~__~__~_ +++

_ 4-

_ +

++ +++

+ + cell body + cell body?d

_

Cat c

_

+++

-

+ +++ (no perikarya) + + endfoot

+ + cell body

-

+ + cell body

++

a Recasens and Delaunoy, 1981. b Inagaki et al., 1985. c Bolz et al., 1985. d Not described clearly.

Inhibitory Nerve Ending

Excitatory Nerve Ending Astrocyte

, k "\ n~a~,'

Ir ..............

[Glutamine

Synthetase

I

~-KG ~

l sAAT l

~(mAAT)

v

GAD

(

Receptor

-'~

(

Receptor

"~

Fig. 10. Synthesis and metabolism of glutamate and GABA in excitatory and inhibitory neurons in combination with astrocytic metabolism. The figure is modified from Kaneko and Mizuno (1994).

glutamine is an energy-saving process. Astrocytes are metabolically coupled with excitatory neurons and thus serve them by supplying precursor glutamine and by de novo synthesizing glutamate from ~-ketoglutarate and ammonia at the cost of ATP and NAD(P)H, respectively. Cortical inhibitory neurons, in contrast to excitatory neurons, appear to use glutamate formed through transamination of ~-ketoglutarate as the immediate precursor of GABA. Since the transamination activity is also an energy-saving process if neurons are supplied with ~-ketoglutarate and aspartate, it may be concluded that neuronal metabolic processes for transmitter synthesis are selected based on the economy of energy as far as possible, and that astrocytes take the part of energy-consuming processes to support the neuronal consumption of transmitter. 226

Enzymes responsible for glutamate synthesis and degradation

Ch. VII

6. ACKNOWLEDGEMENTS

The author is grateful for the photographic help of Mr. Akira Uesugi.

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Norenberg MD (1979): The distribution of glutamine synthetase in the rat central nervous system. J Histochem Cytochem 27:756-762. Norenberg MD, Martinez-Hernandez A (1979): Fine structural localization of glutamine synthetase in astrocytes of rat brain. Brain Res 161:303-310. Recasens M, Delaunoy JP (1981): Immunological properties and immunocytochemical localization of cysteine sulfinate or aspartate aminotransferase-isoenzymes in rat CNS. Brain Res 205:351-361. Reubi JC (1980): Comparative study of the release of glutamate and GABA, newly synthesized from glutamine, in various regions of the central nervous system. Neuroscience 5:2145-2150. Rothe F, Wolf GM, Schtinzel G (1990): Immunohistochemical demonstration of glutamate dehydrogenase in the postnatally developing rat hippocampal formation and cerebellar cortex. Neuroscience 39:419-429. Rothe F, Brosz M, Storm-Mathisen J (1994): Quantitative ultrastructural localization of glutamate dehydrogenase in the rat cerebellar cortex. Neuroscience 62:1133-1146. Rothstein JD, Martin L, Levey AI, Dykes-Hoberg M, Jin L, Wu D, Nash N, Kuncl RW (1994): Localization of neuronal and glial glutamate transporters. Neuron 13:713-725. Schmitt A, Kugler P (1999): Cellular and regional expression of glutamate dehydrogenase in the rat nervous system: non-radioactive in situ hybridization and comparative immunocytochemistry. Neuroscience 92:293-308. Schmitt A, Asan E, Puschel B, Kugler P (1997): Cellular and regional distribution of the glutamate transporter GLAST in the CNS of rats: nonradioactive in situ hybridization and comparative immunocytochemistry. J Neurosci 17:1-10.

Senba E, Kaneko T, Mizuno N, Tohyama M (1991): Somato-, branchio- and viscero-motor neurons contain glutaminase-like immunoreactivity. Brain Res Bull 26:85-97. Shank RE Aprison MH (1981): Present status and significance of the glutamine cycle in neural tissues. Life Sci 28:837-842. Shank RE Campbell GLeM (1983): Metabolic precursors of glutamate and GABA. In: Hertz L, Kvamme E, McGeer EG, Schousboe A (Eds), Glutamine, Glutamate, and GABA in the Central Nervous System. New York: Alan R. Liss, pp 355-369. Shank RE Campbell GLeM (1984a): ~-Ketoglutarate and malate uptake and metabolism by synaptosomes: further evidence for an astrocyte-to-neurons metabolic shuttle. J Neurochem 42:1153-1161. Shank RE Campbell GLeM (1984b): Glutamine, glutamate, and other possible regulators of 0~-ketoglutarate and malate uptake by synaptic terminals. J Neurochem 42:1162-1169. Shapiro RA, Farrell L, Srinivasan M, Curthoys NP (1991): Isolation, characterization, and in vitro expression of a cDNA that encodes the kidney isoenzyme of the mitochondrial glutaminase. J Biol Chem 266:18792-18796. Szerb JC (1984): Storage and release of endogenous and labelled GABA formed from [3H] glutamine and [14C]glucose in hippocampal slices: effect of depolarization. Brain Res 293:293-303. Takatsuna Y, Chiba T, Adachi-Usami E, Kaneko T (1994): Distribution of phosphate-activated glutaminase-like immunoreactivity in the retina of rodents. Curr Eye Res 13:629-637. Tansey FA, Farooq M, Cammer W (1991): Glutamine synthetase in oligodendrocytes and astrocytes: new biochemical and immunocytochemical evidence. J Neurochem 56:266-272. Tapia R, Gonzfilez RM (1978): Glutamine and glutamate as precursors of the releasable pool of GABA in brain cortex slices. Neurosci Lett 10:165-169. Thanki CM, Sugden D, Thomas AJ, Bradford HF (1983): In vivo release from cerebral cortex of [14C]glutamate synthesized from [U-14C]glutamine. J Neurochem 41:611-617. Thompson SG, Wong PT-H, Leong SF, McGeer EG (1985): Regional distribution in rat brain of 1-pyrroline-5-carboxylate dehydrogenase and its localization to specific glial cells. J Neurochem 45:1971-1976. Turman Jr JE, Chandler SH (1994): Immunohistochemical localization of glutamate and glutaminase in guinea pig trigeminal premotoneurons. Brain Res 634:49-61. Ward HK, Thanki CM, Bradford HF (1983): Glutamine and glucose as precursors of transmitter amino acids: ex vivo studies. J Neurochem 40:855-860. Wenthold RJ, Skaggs KK, Altschuler RA (1986): Immunocytochemical localization of aspartate aminotransferase and glutaminase immunoreactivities in the cerebellum. Brain Res 363:371-375. Wenthold RJ, Altschuler RA, Skaggs KK, Reeks KA (1987): Immunocytochemical characterization of glutamate dehydrogenase in the cerebellum of the rat. J Neurochem 48:636-643. Yezierski RE Kaneko T, Miller KE (1993): Glutaminase-like immunoreactivity in rat spinomesencephalic tract cells. Brain Res 624:304-308.

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Sodium- and potassium-dependent excitatory amino acid transporters in brain plasma membranes N.C. DANBOLT

1. INTRODUCTION Glutamate is the major excitatory neurotransmitter in the mammalian nervous system (Fonnum, 1984; Ottersen and Storm-Mathisen, 1984) and exerts its transmitter actions from the extracellular fluid by binding to and thereby activating glutamate receptors. Four families of glutamate receptor proteins (called NMDA, AMPA, kainate and metabotropic receptors) have been identified (for review see: Seeburg, 1993; Dingledine and McBain, 1994; Hollmann and Heinemann, 1994; Schoepfer et al., 1994; Nakanishi et al., 1998; Ozawa et al., 1998). Most neurons and even many glial cells have glutamate receptors in their plasma membranes (for review see: H6sli and H6sli, 1993; Steinhauser and Gallo, 1996; Vernadakis, 1996). Glutamate is involved in most aspects of normal brain function and development (Collingridge and Lester, 1989; Headley and Grillner, 1990; McDonald and Johnston, 1990; LaMantia, 1995), and is thereby both essential and highly toxic (McBean and Roberts, 1985; Choi and Rothman, 1990; Choi, 1992). Consequently, the concentration of glutamate in the extracellular fluid must be tightly controlled. This is not an easy task considering the huge amounts of glutamate (5-15 mmol/kg wet weight depending on the region) in brain tissue (for references see Schousboe, 1981). The highest glutamate concentrations are found inside nerve terminals (Ottersen et al., 1992; Storm-Mathisen et al., 1992). The extracellular concentration of glutamate is normally in the low micromolar range (Hamberger et al., 1983). The only (significant) mechanism capable of removing glutamate from the extracellular fluid is the glutamate uptake system which detoxifies glutamate by pumping it into cells (Balcar and Johnston, 1972; Logan and Snyder, 1972; Johnston, 1981). This uptake system consists of a family of Na +- and K+-coupled glutamate transporters (for review see: Kanai et al., 1993, 1997; Danbolt, 1994, 1998b; Gegelashvili and Schousboe, 1997; Robinson and Dowd, 1997). The roles of glutamate transporters for brain physiology are not fully understood. It is clear that the transporters play important roles in glutamate removal and that this is essential both for securing a high signal-to-noise ratio in glutamatergic transmission and for avoiding excitotoxicity (harmful glutamate receptor overactivation). These tasks appear relatively simple. The glutamate uptake system, however, is complex and consists of several different transporter proteins which have highly differentiated and dynamically regulated localizations. The transport activities and transporter concentrations are also subject to regulation. The transporters even have chloride-channel activities. Thus, this is a sophisticated system which probably plays more Handbook of Chemical Neuroanatomy, Vol. 18: Glutamate O.P. Ottersen and J. Storm-Mathisen, editors (D 2000 Elsevier Science B.V. All rights reserved.

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refined roles than simple transmitter removal. The transporters may modify the time course of synaptic events, the extent and pattern of activation of receptors outside the synaptic cleft and at neighboring synapses (intersynaptic crosstalk) as well as the level of receptor desensitization (for review see: Beckman and Quick, 1998; Danbolt et al., 1998b; Bergles et al., 1999; Hediger, 1999; Kullmann, 1999; Seal and Amara, 1999; Sims and Robinson, 1999).

2. GLUTAMATE TRANSPORTER TYPES A variety of glutamate transporters exist in the brain (for review see Danbolt, 1994). These include intracellular glutamate transporters that carry dicarboxylic amino acids across the inner mitochondrial membrane and the as yet molecularly unidentified transporter(s) which loads synaptic vesicles with glutamate from the cytoplasm. The plasma membranes contain the so-called 'sodium-dependent high-affinity' glutamate transporters, which are the topic of this review, and a variety of other transporters, including the glutamate-cystine exchangers (Sato et al., 1999) as well as a number of poorly characterized transporters like the glutamate-ascorbate exchanger, the glutamate-GABA exchanger and others (for review see Danbolt, 1994). The 'sodium-dependent high-affinity' glutamate transporters will in the rest of this review simply be referred to as 'glutamate transporters'. Five such glutamate transporters have been cloned so far (for review see: Saier, 1999; Slotboom et al., 1999): GLAST (EAAT1) (Storck et al., 1992; Tanaka, 1993), GLT (EAAT2) (Pines et al., 1992), EAAC (EAAT3) (Kanai and Hediger, 1992), EAAT4 (Fairman et al., 1995) and EAAT5 (Arriza et al., 1997). 1 In addition to these five cloned glutamate transporters, there is another one, namely the one in the plasma membranes of glutamatergic nerve endings (Divac et al., 1977; Storm-Mathisen, 1977; Fonnum, 1984). This transporter has not been identified by molecular cloning and it is not recognized by any of the antibodies so far made to known glutamate transporters (Danbolt e,t al., 1998b). The strongest evidence yet presented for the existence of this transporter is the electron microscopic immunocytochemistry with antibodies to D-aspartate (Gundersen et al., 1993, 1996). In these studies, brain tissue slices were pre-incubated (45 rain, 30~ in oxygenated Krebs' solution), incubated (20 min) with 10 or 50 IxM D-aspartate, fixed in glutaraldehyde and subjected to quantitative postembedding electron microscopical immunocytochemistry using antibodies to glutaraldehyde-fixed o-aspartate and gold-particle-tagged secondary antibodies. A sodium-dependent, threo-hydroxyaspartate-sensitive accumulation of D-aspartate immunoreactivity was detected in nerve terminals (implying millimolar concentrations of D-aspartate as judged from simultaneously processed test sections with known amounts of fixed amino acid). Because O-aspartate is very slowly metabolized in the adult brain (Davies and Johnston, 1976; Takagaki, 1978), it is hard to explain how D-aspartate can be concentrated about two orders of magnitude in the terminals within 20 min unless it is taken up by a glutamate transporter.

3. MECHANISM OF GLUTAMATE UPTAKE The uptake process is driven by the electrochemical gradients across the cell membrane. Sodium is required for glutamate binding while potassium is required for net transport I The actual meanings of the acronyms (GLAST, glutamate-aspartate transporter; GLT, glutamate transporter; EAAC, excitatory amino acid carrier; EAAT, excitatory amino acid transporter) are not important, as they do not reflect functional differences among the transporters.

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(Kanner and Sharon, 1978; Sarantis and Attwell, 1990). The transporters utilize the gradients of Na +, K + and H + as energy sources for the transport process. The stoichiometry of the process is an important parameter because it determines the concentrative capacity, the energy consumption and the sensitivity of the transport process to ischemia and other perturbations of the driving forces. According to recent studies (Zerangue and Kavanaugh, 1996; Levy et al., 1998), the EAAC and GLT glutamate transporters have the following stoichiometry: 1 glutamate is taken up together with 3 Na + and 1 H + in exchange for 1 K +. In addition to being co-transporters, the glutamate transporter proteins also function as chloride channels (Sonders and Amara, 1996; Fairman and Amara, 1999; Seal and Amara, 1999). The chloride conductance is triggered by sodium-dependent glutamate binding, but is thermodynamically independent of the transport process. Consequently, the transporters behave as glutamate-gated chloride channels. This property is particularly prominent in EAAT4 and EAAT5 and almost non-existent in GLT (Fairman et al., 1995; Vandenberg et al., 1995; Wadiche et al., 1995a,b; Arriza et al., 1997). EAAT4 also has a proton (H +) conductance that is controlled by glutamate and arachidonic acid (Fairman et al., 1998). Thus, the glutamate transporters are not simple transporter molecules. EAAT4 and EAAT5 may to some extent function as inhibitory glutamate receptors because of the high chloride conductance.

4. L O C A L I Z A T I O N OF GLUTAMATE T R A N S P O R T E R S To understand glutamatergic neurotransmission it is necessary to obtain precise information on the localizations and densities of glutamate transporters because they represent one of the major determinants of how glutamate diffuses from the point of release (Fig. 1). Since the cloning of the first glutamate transporters in 1992, a substantial amount of information on transporter distribution has been collected. Unfortunately, the literature is starting to become somewhat confusing because clear distinction is not always made between transporter protein and transporter mRNA and between cells in culture and cells in the intact brain. Further, antibodies to different epitopes on the same protein may give different results if the protein is subject to variable splicing or if one of the epitopes is masked or partially masked by an interacting protein. For this reason, information on the epitopes recognized by the antibodies may be important. This information is known whenever synthetic peptides have been used to generate the antibodies and should be stated in the publications where the antibodies are used. Finally, the importance of proper testing of the specificity of antibodies for immunocytochemistry cannot be overemphasized (for a detailed discussion see Danbolt et al., 1998a). 4.1. LOCALIZATION OF GLT (EAAT2)

4.1.1. GLT is the major glutamate transporter in the forebrain The most abundant glutamate transporter in the mammalian forebrain is GLT and it dominates in all the regions of the central nervous system except those few where GLAST is the major transporter (see below). The quantitative importance of GLT in the forebrain is apparent from several different studies. Firstly, it was GLT that was isolated when transport activity was used to monitor the purification process (Danbolt et al., 1990; Pines et al., 1992). Secondly, most of the reconstitutable transport activity in crude detergent extracts of forebrain tissue can be immunoprecipitated with GLT antibodies (Danbolt et al., 1992; Haugeto et al., 1996). Thirdly, 233

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Fig. 1. Schematic representation of the localization of glutamate transporters in the vicinity of the glutamatergic parallel fiber-to-Purkinje cell dendritic spine synapses in the cerebellar molecular layer. GLAST and GLT are intermingled in the astrocytic membranes at average densities of 4700 and 740 I~m-2 membrane, respectively (Lehre and Danbolt, 1998). The concentrations are highest near synapses and are lower along major dendrites. EAAT4 is concentrated in the glia-covered parts of the membranes of Purkinje cell dendrites, highest at the spines and thinner dendrites. There is a very low concentration of EAAT4 at the postsynaptic density. EAAT4 is unevenly distributed in the molecular layer and is expressed at different densities in different parasagittal zones, average density at around 1800 molecules itm -2 (Dehnes et al., 1998). EAAC is present in the Purkinje cell plasma membrane as well as cytoplasm (Conti et al., 1998; Kugler and Schmitt, 1999). Quantitative data and information on the precise subcellular distribution are currently unavailable. Two (or more) glutamate transporters remain to be identified by molecular cloning, namely the one in glutamatergic nerve terminals and the one in the synaptic vesicles (both marked in red).

mutant mice lacking GLT show lethal spontaneous seizures and increased susceptibility to acute forebrain injury (K. Tanaka et al., 1997). Brain tissue homogenates from these animals have m u c h lower glutamate uptake activity than similar homogenates from wild-type mice. Mice lacking G L A S T have increased susceptibility to cerebellar injury as well as reduced motor coordination (Watase et al., 1998), while mice deficient in E A A C (Peghini et al., 1997) develop behavioral abnormalities, but no neurodegeneration. Fourthly, m e a s u r e m e n t s of transporter protein have shown that the level of GLT is about 4 times higher than G L A S T in the hippocampus and 1/6 in the cerebellum (Lehre and Danbolt, 1998).

4.1.2. Exclusive glial expression of GLT protein, but not of GLT mRNA GLT protein has so far been detected exclusively in astroglial cells in the normal adult central nervous system (excluding retina; see Section 4.1.3 below). S o m e investigators (e.g. 234

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Schmitt et al., 1996) who have not employed electron microscopy, have expressed themselves cautiously since they have not been able to make accurate distinctions between neuronal elements and closely associated glial processes, but no one has so far reported neuronal labeling. In an initial study (Chaudhry et al., 1995), we noted a weak GLT immunoreactivity at around background levels in some hippocampal nerve terminals. We later followed this up, but neuronal GLT labeling statistically significantly different from background could not be demonstrated (F.A. Chaudhry and J. Storm-Mathisen, unpublished). Thus, the GLT protein has so far only been found in astrocytes (fibrous as well as protoplasmic) in the normal adult rat brain and spinal cord (Danbolt et al., 1992; Hees et al., 1992; Levy et al., 1993b; Rothstein et al., 1994; Chaudhry et al., 1995; Lehre et al., 1995; Schmitt et al., 1996) as well as in the adult human brain (Milton et al., 1997). This includes fibrous astrocytes in white matter. No astrocyte in the hippocampus or in the cerebellum has been identified as GLT-deficient (Chaudhry et al., 1995). Further, GLT protein has neither been found in oligodendrocytes nor in epithelial cells of the choroid plexus or in tanycytes, but possibly in activated microglia (Swanson et al., 1997; Lopez-Redondo et al., 1999). In agreement with the localization of the protein, the mRNA encoding GLT is expressed in astroglial cells (Torp et al., 1994, 1997; Schmitt et al., 1996; Berger and Hediger, 1998). However, the mRNA is also present in the majority of the neurons in the neocortex and also in the olfactory bulb and in pyramidal cells in CA3 hippocampus (Torp et al., 1994, 1997; Schmitt et al., 1996; Berger and Hediger, 1998). The reason why neurons do not normally express GLT in spite of the fact that they produce the mRNA, is not known. Perhaps the explanation is to be found in the length of the mRNA molecule which is 11.3 kb or 6.6 times longer than the coding sequence (Pines et al., 1992). The antibodies used to detect GLT in tissue sections bind to epitopes on the N-terminal first 34 residues or to epitopes on the last (C-terminal) 80 residues. It therefore cannot be excluded that a novel GLT variant lacking these epitopes is present in neurons. Alternatively, the neuronal GLT expression must be orders of magnitude lower than that of astrocytes. 4.1.3. GLT protein in neurons

Although GLT has only been detected in astroglial cells in the normal and mature nervous system [with the exception of retina where bipolar cells and amacrine cells normally express GLT protein (Rauen et al., 1996)], this does not mean that neurons never express GLT. Several populations of neurons express GLT during the development of the nervous system (see Section 4.6), but the neuronal expression is transient and disappears on maturation. GLT has also been frequently observed in cultured neurons (Brooks-Kayal et al., 1998; Mennerick et al., 1998; Meaney et al., 1998; Stanimirovic et al., 1999; Plachez et al., 2000). In newborn piglets, GLT may also appear in neurons after hypoxia-ischemia (Martin et al., 1997) showing that the cellular expression can potentially change. 4.1.4. Regional and subcellular distribution of GLT in adult rat brain tissue

The highest GLT levels are found in the forebrain. Compared to hippocampus, the concentrations of GLT protein in cerebral cortex, thalamus, bulbus olfactorius and cerebellum are 93, 54, 30 and 24%, respectively (Lehre et al., 1995; 12t. Haugeto and N.C. Danbolt, unpublished). GLT, as well as GLAST (see Section 4.2) and EAAT4 (see Section 4.4), are normally predominantly present at the surface of cells in the brain (Chaudhry et al., 1995; Lehre et al., 235

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1995). The immunoreactivity of membranes in the cytoplasm is very low and at background levels compared to that of the plasma membrane (Chaudhry et al., 1995). With quantitative (postembedding) immunocytochemistry and freeze-substituted and lowtemperature embedded tissue, it has been shown (Chaudhry et al., 1995) that the concentrations of GLT (as well as GLAST) in the membranes not only differ between astrocytes from different regions, but that the concentrations also differ between different parts of the same cell. The variations correlate with the type of neighboring structure. No concentration differences were found between the cell bodies and the processes. Astroglial membranes facing neuropil have higher densities than astroglial membranes facing other astrocytes, cell bodies, pia mater or capillary endothelium (Chaudhry et al., 1995). By using known amounts of pure GLT protein (Lehre and Danbolt, 1998) as standard during quantitative immunoblotting (Levy et al., 1995), it has been possible to determine the concentrations of GLT and GLAST in the hippocampus and in the cerebellar molecular layer in absolute terms. By combining these data with data on the plasma membrane areas (Lehre and Danbolt, 1998), it was concluded that the concentration of GLT in hippocampus (stratum radiaturn, CA1) is as high as 12,000 molecules per ixm3 tissue or 8500 molecules per i~m2 astroglial cell membrane, while the concentration in the cerebellar molecular layer is considerably lower (2800 txm -3, 740 txm -2) (Lehre and Danbolt, 1998). It should be noted that these values are from adult rat brain, that the concentrations change during development (see Section 4.6) and that the expression is regulated via a number of different mechanisms (see Section 5). 4.2. LOCALIZATION OF GLAST (EAAT1) Like GLT, GLAST is expressed throughout the CNS, but at different concentrations in different regions (Lehre et al., 1995; Schmitt et al., 1997; Berger and Hediger, 1998). GLAST is more abundant than GLT in the cerebellum (Lehre and Danbolt, 1998), the inner ear (Furness and Lehre, 1997; Takumi et al., 1997), the circumventricular organs (Berger et al., 2000) and in the retina (Derouiche and Rauen, 1995; Derouiche, 1996; Rauen et al., 1996, 1998; Lehre et al., 1997; Pow and Barnett, 1999). Results of studies of mice lacking GLAST fit nicely with these data. The GLAST-deficient mice show symptoms of insufficient glutamate uptake in the cerebellum, namely increased susceptibility to cerebellar injury as well as reduced motor coordination (Watase et al., 1998) and have major changes in the retina, abnormal electroretinogram and exacerbated damage after ischemia (Harada et al., 1998). The amount of GLAST is about 6 times higher than that of GLT in the cerebellum (Lehre and Danbolt, 1998). 4.2.1. Cellular distribution of GLAST in the CNS

The localization of GLAST is more straightforward than that of GLT: both GLAST protein (Chaudhry et al., 1995; Lehre et al., 1995; Schmitt et al., 1997) and GLAST mRNA (Torp et al., 1994; Schmitt et al., 1997; Berger and Hediger, 1998) are expressed by astroglial cells throughout the entire CNS. No astrocytes have so far been identified as GLAST-deficient. GLAST and GLT are expressed in the same astrocytes (Lehre et al., 1995; Haugeto et al., 1996), but in different proportions in different parts of the brain (Lehre et al., 1995) and co-exist in the same astroglial cell membranes as separate homo-oligomeric complexes (Haugeto et al., 1996; Kavanaugh, 1999). GLAST mRNA and protein are found in high concentrations close to the ventricles in a subependymal glial plexus (Torp et al., 1994; Lehre et al., 1995), but in lower concentrations also in the ependymal cells (Schmitt et al., 1997; Berger and Hediger, 1998). 236

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Neuronal expression of GLAST protein has not been detected anywhere in the nervous system in vivo, neither in the adult nor during development, although a possible expression of GLAST mRNA in granule cells of the mouse hippocampus (gyms dentatus) at birth (P0) has been described (Sutherland et al., 1996), but this has not been verified with antibodies. However, it has recently been reported that GLAST protein is transiently expressed in a subpopulation of cultured embryonic hippocampal neurons (Plachez et al., 2000). The neuronal expression of GLAST initially reported (Rothstein et al., 1994) does not count, as the authors themselves report (Ginsberg et al., 1995; Rothstein et al., 1995) that the antibody used was not good enough and that GLAST is expressed only in astroglial cells, in agreement with reports by other groups (Chaudhry et al., 1995; Lehre et al., 1995; Schmitt et al., 1997). The 'rule' applies to retina as well: GLAST is present in both Mtiller cells and in regular astrocytes (Derouiche and Rauen, 1995; Derouiche, 1996; Rauen et al., 1996, 1998; Lehre et al., 1997). It has been debated if GLAST is expressed in the retinal pigment epithelium and in oligodendrocytes. Expression has been reported by some (Derouiche and Rauen, 1995; Choi and Chiu, 1997; Domercq and Matute, 1999; Domercq et al., 1999), but questioned by others (Lehre et al., 1995, 1997; Schmitt et al., 1997). Finally, GLAST is present in fibrocytes and in supporting cells of inner ear (Furness and Lehre, 1997; Takumi et al., 1997). 4.2.2. Subcellular distribution of GLAST Like GLT and EAAT4, most of the GLAST protein is normally (adult rat brain) found in the plasma membranes. Very little is seen in the cytoplasm (Chaudhry et al., 1995; Lehre et al., 1995). A similar picture is observed in the inner ear (Furness and Lehre, 1997; Takumi et al., 1997). No differences in GLAST densities have been noted between cell bodies and processes (Chaudhry et al., 1995). The observed variations in GLAST densities within individual astrocytes correlate with the type of neighboring structure in the same way as is described above for GLT (Chaudhry et al., 1995). Astrocytic membranes facing neuropil have higher densities than membranes facing capillary endothelium, cell bodies, large dendrites and pia mater. This highly differentiated localization probably implies that the transporters are kept in the correct positions in the membrane by other proteins. A careful regulation of GLAST expression and targeting is also suggested. 4.2.3. Concentrations of GLAST protein The highest concentrations of GLAST are seen in the molecular layer of the cerebellum, the Bergmann glia in particular (Lehre et al., 1995). Compared to the cerebellum, the concentrations of GLAST protein in bulbus olfactorius, hippocampus, cerebral cortex and thalamus are 49, 35, 33 and 22%, respectively (Lehre et al., 1995; Lehre and Danbolt, 1998; O. Haugeto and N.C. Danbolt, unpublished). The density of GLAST in the molecular layer is as high as 18,000 GLAST molecules per i~m3 tissue (molecular layer) or about 4700 molecules per ixm2 Bergmann glia cell membrane (Lehre and Danbolt, 1998). The concentration of GLAST in the hippocampus (stratum radiatum, CA1) is 3200 molecules per i~m3 tissue or about 2300 per i~m2 astroglial cell membrane. 237

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4.3. LOCALIZATION OF EAAC (EAAT3) 4.3.1. Antibodies to EAAC

Less information is available on the localization of EAAC than on GLT, GLAST and EAAT4. The reasons for this are (a) that the concentration of EAAC is considerably lower than those of the others (Haugeto et al., 1996), making it more difficult to detect on immunoblots and on ultra thin sections (postembedding for quantitative electron microscopy), and (b) that it has been more difficult to find out if the antibodies are specific. EAAC has a confusing behavior on immunoblots, binds poorly to nitrocellulose and PVDF membranes (Y. Dehnes and N.C. Danbolt, unpublished) and is present in significant amounts in the cytoplasm (Conti et al., 1998; Kugler and Schmitt, 1999). It is now clear that the cytoplasmic immunoreactivity is real and due to a rapidly mobilizable pool of transporter proteins (see Section 5) rather than a sign of poor antibody specificity. The majority of the published studies on the distribution of EAAC are based on antibodies (Rothstein et al., 1994), to the C-terminal 14 amino acid residues (511-524) of EAAC (Kanai and Hediger, 1992), a sequence that is identical in rabbit, mouse, rat and man (BjCrgts et al., 1996). Antibodies have also been produced to residues 480-499 of rat EAAC (Kugler and Schmitt, 1999), and to residues 510-524 (Haugeto et al., 1996) and 491-524 (Y. Dehnes and N.C Danbolt, unpublished) of rabbit EAAC. In view of the difficulties in proving monospecificity of the EAAC antibodies (see above), it is surprising that none of the published reports show immunoblots containing all tissue antigens (for discussion of antibody testing see Danbolt et al., 1998a). Nevertheless, the results obtained with Rothstein's and Kugler's antibodies seem to be in agreement with each other and with our own (preliminary) results (Y. Dehnes, K. Ullensvang, K.R Lehre and N.C. Danbolt, unpublished). 4.3.2. Localization of EAAC in the adult CNS

EAAC is widely distributed in the body. It is strongly expressed in peripheral organs, kidneys and small intestine in particular. Within the CNS, the highest levels are found in the hippocampus, cerebellum and basal ganglia (Rothstein et al., 1994; Conti et al., 1998; Kugler and Schmitt, 1999). Based on immunoblots, it is believed that the concentration of EAAC is lower than those of the GLT, GLAST and EAAC (Haugeto et al., 1996), but the quantitative data are unreliable due to the poor binding to blotting membranes (see above). However, the notion that EAAC is expressed at lower levels than the other mentioned transporters, is supported by the studies of EAAC-deficient mice, which display kidney and behavioral abnormalities, but no neurodegeneration (Peghini et al., 1997). EAAC mRNA (Kanai and Hediger, 1992) and EAAC protein (Rothstein et al., 1994) are expressed in neurons in the rat brain. In fact, EAAC is present in several types of neurons, including GABAergic neurons. Importantly, the labeling is concentrated in the neuronal cell bodies (somata) and dendrites apparently avoiding the nerve terminals (Rothstein et al., 1994). EAAC mRNA and protein are both present in most if not all glutamatergic neurons, as well as in several GABAergic and cholinergic neurons (Meister et al., 1993; Kanai et al., 1995; BjCrfis et al., 1996; Velaz-Faircloth et al., 1996; Torp et al., 1997; Berger and Hediger, 1998; Conti et al., 1998; Kugler and Schmitt, 1999). EAAC is also present in astrocytes of the cerebral cortex and white matter (Conti et al., 1998) as well as in oligodendrocytes in various white matter regions (Domercq and Matute, 1999; Domercq et al., 1999; Kugler and Schmitt, 1999). Further, it is also found (Kugler and Schmitt, 1999) in peripheral neurons (spinal ganglia) and 238

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in oligodendrocytes in various white matter regions of the CNS, in ependymal cells, and in epithelial cells of the plexus choroideus of the four ventricles, as well as in satellite cells of spinal ganglia. In the retina, EAAC is located in horizontal cells, amacrine cells, displaced amacrine cells, ganglion cells as well as in some bipolar cells, but not in Mtiller cells (Rauen et al., 1996; Schultz and Stell, 1996). A striking feature of the EAAC immunoreactivity is that a large part of it localizes to cytoplasmic structures (Conti et al., 1998; Kugler and Schmitt, 1999). The cytoplasmic localization of EAAC immunoreactivity is in sharp contrast to the predominant plasma membrane localizations of GLT, GLAST (Chaudhry et al., 1995) and EAAT4 (Dehnes et al., 1998). It is now believed to imply that EAAC can be rapidly mobilized from the cytoplasm to the plasma membrane (see Section 5). EAAC is, at least in the cerebral cortex, present throughout the dendritic ramifications, including the spines and is therefore close to the synapses (Conti et al., 1998). The pre-embedding technique used in the latter study enabled the authors to conclude that the spines contain EAAC, but not which parts of the spine membrane that contains EAAC (for review of the methods see Danbolt et al., 1998a). Thus, it is not known if EAAC is present only in the non-synaptic parts of the spine membrane or if it is also present in the synaptic area. Further, it is difficult to assess the importance of EAAC before information on the concentration becomes available. 4.4. LOCALIZATION OF EAAT4 4.4.1. Regional and cellular distribution of EAAT4 EAAT4 has only been detected in one cell type in the adult rat CNS, namely the Purkinje cells of the cerebellar molecular layer (Yamada et al., 1996; Nagao et al., 1997; J. Tanaka et al., 1997; Dehnes et al., 1998). The localization is the same in man (Bar-Peled et al., 1997; Furuta et al., 1997a; Itoh et al., 1997; Inage et al., 1998). Although EAAT4 is mainly expressed in the cerebellum, there is some EAAT4 in the forebrain too. The concentration is very low, but it has been possible to isolate EAAT4 from adult rat forebrain by means of antibodies (Dehnes et al., 1998). Some proteins, e.g. zebrin (for review see Hawkes, 1997), are expressed in the cerebellar molecular layer in a zonal pattern. Thus, some cells contain relatively high concentrations of zebrin, while others contain much less. It turns out that the expression of EAAT4, in contrast to the other glutamate transporters, follows that of zebrin. The Purkinje cells with high zebrin levels also have high EAAT4 levels (Nagao et al., 1997; Dehnes et al., 1998). The functional significance of these zones is unknown, but has been reported to correspond to tactile projection patterns (Hallem et al., 1999). 4.4.2. Subcellular distribution in the adult Purkinje cells There are not only differences in EAAT4 densities among different Purkinje cells, but also within individual cells. Almost all the EAAT4 is found in the plasma membranes of the cell bodies and dendrites, including the spines. There are low levels in cytoplasmic structures, with the exception of multivesicular bodies (Yamada et al., 1996; Furuta et al., 1997a; Itoh et al., 1997; Dehnes et al., 1998). The concentration of EAAT4 is highest in the spine membranes and drops gradually towards the cell bodies. The expression is low in the synaptic area (J. Tanaka et al., 1997), but it is not zero (Dehnes et al., 1998). No signal was detected in 239

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the perisynaptic membrane. Interestingly, EAAT4 is virtually only expressed in the parts of the spine-membrane-facing astroglia. The average concentration of EAAT4 in the molecular layer of the adult rat is about 1900 molecules per i~m3 tissue. Because the surface area of the spines is about 1.1 txmZ/~m 3 (Dehnes et al., 1998), it follows that the average density of EAAT4 is about 1800 transporter molecules per i~m2 spine membrane (Dehnes et al., 1998). 4.5. LOCALIZATION OF EAAT5 Two variants of EAAT5 (sEAAT5A and sEAAT5B) have been isolated from salamander retina (Eliasof et al., 1998a) and are expressed in both MUller cells and in most of the neurons (Eliasof et al., 1998a,b), but there are as yet no publications describing the localization of EAAT5 in mammals, except from Northern blotting showing a strong signal in retina and no detectable signal in the brain (Arriza et al., 1997). 4.6. DEVELOPMENTAL CHANGES IN GLUTAMATE TRANSPORTER EXPRESSIONS As mentioned above, glutamate-mediated signaling is important in the regulation of the nervous system development (for review see: McDonald and Johnston, 1990; Komuro and Rakic, 1993; Johnston, 1995; Vallano, 1998). There are dynamic changes in the expression and subunit composition of the NMDA receptors (for review see Watanabe, 1997) as well as in that of AMPA, kainate and metabotropic receptors (e.g. Ryo et al., 1993; Bahn et al., 1994; Catania et al., 1994; Jakowec et al., 1995; Minakami et al., 1995; Romano et al., 1996; Paschen et al., 1997). Glutamate modulates neuronal migration (Komuro and Rakic, 1993; Rossi and Slater, 1993), and is important for synapse elimination (Rabacchi et al., 1992). In line with this, both overstimulation (Johnston, 1995) and blockade (Deutsch et al., 1998) of glutamate receptors are harmful to the developing brain. In view of the importance of glutamate for the development of the nervous system, it seems important to gain information on the glutamate transporter during development.

4.6.1. Changes in transporter concentrations It has been known for some time that brain glutamate uptake activity is low at early developmental stages and that it increases sharply at around the most active period of synaptogenesis (Schousboe et al., 1976; Schmidt and Wolf, 1988; Kish et al., 1989; Erd6 and Wolff, 1990; Christensen and Fonnum, 1992; Collard et al., 1993; Cohen and Nadler, 1997). In line with this, the concentrations of both GLT and GLAST proteins (in the rat) are present from early developmental stages at low concentration and increase dramatically in the most active period of synaptogenesis (from the end of the second postnatal week and to the end of the fourth week) reaching near adult levels by P35 (Furuta et al., 1997b; Ullensvang et al., 1997). The highest levels of GLT and GLAST mRNA are observed at P14 in the mouse forebrain (Shibata et al., 1996; Sutherland et al., 1996). The increase in reconstitutable transport activity (from rat forebrain) parallels that of the GLT protein expression (Ullensvang et al., 1997) in agreement with the notion that GLT is the major glutamate transporter (see Section 4.1.1). The changes in GLT levels are more dramatic than the changes in GLAST levels as GLAST is easily detectable at birth while GLT is not. The concentration of GLT then increases so much that it becomes higher than that of GLAST. In the cerebellum, the largest increase in GLT is observed between P21 and P35 (Ullensvang et al., 1997). 240

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The protein levels of EAAC and EAAT4 also display major changes. The highest concentration of EAAC is seen at around postnatal day 5 (P5) in the rat and gestational day 60 in sheep before it declines to adult levels (Furuta et al., 1997b; Northington et al., 1998). EAAT4, which has not been reported before birth, is present in rat cerebellum at low concentrations at P1 and increases strongly from P10 onwards to reach adult levels about two weeks later (Furuta et al., 1997b). The developmental pattern of expression seems to be similar in man (Itoh et al., 1997). Interestingly, EAAT4 is transiently expressed at quite high levels in the rat forebrain where it reaches a maximum at around P10 before it declines to adult levels (Furuta et al., 1997b; C. Plachez and N.C. Danbolt, unpublished). The localization of the protein in the forebrain during its transient expression has not been studied. 4.6.2. Changes in the localizations of GLT and GLAST In two elegant studies on mouse spinal cord (Shibata et al., 1997; Yamada et al., 1998) Watanabe's team show that GLT is expressed in differentiating neurons, while GLAST is found on the directional cellular elements along which young neurons elongate their axons or move their cell bodies, namely the radial glia. The cell bodies of the radial glia are located in the ventricular or subventricular zone and have long slender radially oriented processes penetrating the marginal zone and ending on the pial surface. Radial glia comprises a distinct class of neuroglia that guides neurons during their migration and that later transforms into astrocytes and oligodendrocytes (Rakic, 1971; Choi, 1981; Raft et al., 1983). Thus, at E l l and E 13 GLAST mRNA is found in the ventricular zone while GLAST immunoreactivity (protein) is found in the cell bodies as well as in the radially oriented processes that extend from the ventricular zone, through the marginal zone and end on the glial surface. On the other hand, GLT (protein and mRNA) is found in a different population of cells (interpreted as neurons) in the marginal zone at El3. GLT immunoreactivity is not detected in the radial fibers. At P7, however, the neuronal GLT immunoreactivity has disappeared and GLT protein is now co-localized with GLAST in astrocytes. This fits with other studies showing an exclusive astroglial localization of GLAST and GLT postnatally in rodents (Furuta et al., 1997b; Ullensvang et al., 1997). Electron microscopical immunocytochemistry of rat hippocampus (Ullensvang et al., 1997) reveals exclusive astroglial localization from the moment GLAST and GLT become detectable with the method used (P6 and P 1 l, respectively). Transient neuronal expression of GLT has also been observed in sheep (Northington et al., 1998, 1999).

5. REGULATION OF GLUTAMATE UPTAKE Glutamate uptake seems to be under regulatory control on virtually all possible levels, i.e. DNA transcription, mRNA splicing, protein synthesis, protein targeting, and amino acid transport and associated ion-channel activities (for review see: Gegelashvili and Schousboe, 1998; Sims and Robinson, 1999; Gegelashvili et al., 2000). 5.1. GLUTAMATE TRANSPORTER EXPRESSION Lesioning of glutamatergic fibers leads to a reduction in glutamate uptake activity in the target area of the lesioned fibers (Divac et al., 1977; Storm-Mathisen, 1977; Fonnum, 1984). This is not only due to a loss of nerve terminals and thereby to a loss of nerve terminal glutamate transporters as originally believed (for review see: Fonnum, 1984; Ottersen and 241

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Storm-Mathisen, 1984), but also to a loss of glial glutamate transporters (GLT and GLAST) in the target area. Thus, lesions of the rat cerebral cortex lead to a reduction in GLT and GLAST as well as synaptosomal glutamate uptake in the striatum 7 days after the operation (Levy et al., 1993a, 1995; Ginsberg et al., 1995), but not in EAAC (Ginsberg et al., 1995). These findings strongly suggest that neurons influence the transporter expression in glial cells. This has now been shown more directly. It appears that the glial expression of glutamate transporters depends on soluble factors released from neurons (for review see: Gegelashvili and Schousboe, 1998; Sims and Robinson, 1999; Gegelashvili et al., 2000). Astrocytes cultured in the absence of neurons express GLAST, but very little GLT (Kondo et al., 1995; Gegelashvili et al., 1996). The expression of GLT is turned on (or strongly up-regulated) when the astrocytes are grown together with neurons (co-cultures) or grown in neuron-conditioned medium (no cells) (Gegelashvili et al., 1997; Swanson et al., 1997; Schlag et al., 1998). The nature of the stimulatory factor(s) in the neuron-conditioned media is still unknown. The astrocytic expression of GLAST, as well as the D-aspartate uptake activity, are up-regulated upon stimulation of the kainate-preferring type of glutamate receptors (Gegelashvili et al., 1996). In contrast, glutamate receptors do not up-regulate GLT (Gegelashvili et al., 1997). 5.2. POSTTRANSLATIONAL REGULATION OF TRANSPORTERS Posttranslational regulation of glutamate transporters includes changes in cell surface expression due to trafficking between the plasma membrane and intracellular compartments, and modulation of transport activity by direct phosphorylation, redox modulation of sulfhydryl groups and inhibition by arachidonic acid and other cis-polyunsaturated fatty acids. The activities of some membrane proteins can be regulated by adding them to or removing them from the cell surface. This mechanism allows rapid changes without having to synthesize new protein. The most famous example is perhaps the glucose transporter GLUT4 which moves to the plasma membrane in response to insulin (for review see: Rea and James, 1997; Pessin et al., 1999). But also the transporters for GABA and dopamine are regulated by this mechanism (Corey et al., 1994; Quick et al., 1997; Zhu et al., 1997; Bernstein and Quick, 1999; Melikian and Buckley, 1999). Now, rapid changes in the cell surface expression of EAAC (Davis et al., 1998), GLAST (Duan et al., 1999) and EAAT4 (Gegelashvili et al., 2000) have been observed in C6 glioma cells, primary cultures of murine astrocytes and BT4C glioma cells, respectively. The trafficking of EAAC is controlled by protein kinase-C-mediated phosphorylation, while the signal triggering the increases in surface expression of GLAST and EAAT4 seems to be the transporter substrate (e.g. glutamate and aspartate). Arachidonic acid inhibits several sodium-coupled amino acid transporters including the uptake systems for glutamate, glycine and GABA (Chan et al., 1983; Rhoads et al., 1983; Yu et al., 1986; Barbour et al., 1989; Zafra et al., 1990; Volterra et al., 1992, 1994; Lynch et al., 1994; Lundy and McBean, 1995; Breukel et al., 1997; Manzoni and Mennini, 1997). This effect of arachidonic acid (Volterra et al., 1994; Trotti et al., 1995) is distinct from and additive to the effects of oxidation (see below) and independent of the effects of arachidonic acid on the electrochemical gradients across the cell membranes. Studies on purified and reconstituted rat brain GLT (Trotti et al., 1995) confirm that arachidonic acid inhibits this transporter. The inhibitory effect is due to direct action on the transporter itself rather than an effect through other mechanisms or via the phospholipid membrane. Further, the arachidonic acid ethyl ester is inactive, suggesting that the free carboxylic group is required for inhibitory activity. Human GLAST expressed in Xenopus oocytes is also inhibited by arachidonic acid, human GLT is 242

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stimulated (Zerangue et al., 1995). Arachidonic acid also activates a glutamate-gated proton (H +) conductance in EAAT4 (Fairman et al., 1998; Tzingounis et al., 1998). It turns out that the transport activity of GLAST, GLT and EAAC (individually expressed in HeLa cells, solubilized and reconstituted in liposomes) can be turned down and up by sequential treatment with 5,5'-dithio-bis(2-nitrobenzoic) acid (DTNB, a thiol-oxidizing agent) and DTT, respectively. These data suggest that the glutamate transporters possess an SH-based redox regulatory mechanism (Trotti et al., 1997b). The redox interconversion of SH groups on EAAC-reduced Vmaxof glutamate transport without affecting Km and without affecting the C1--conductance (Trotti et al., 1997a). Direct phosphorylation of the glutamate transporter proteins themselves has only been reported for GLT and GLAST. Protein kinase C phosphorylates GLAST and thereby reduces the transport activity to 25% with no change in cell surface expression (Conradt and Stoffel, 1997). GLT was originally reported to be stimulated by phosphorylation of serine-113 (Casado et al., 1993), but a recent report (Tan et al., 1999) suggests that the protein kinase-C-mediated stimulation represents an effect of the expression system used rather than an effect on GLT. Further studies are required to sort out the controversy.

6. T H E R O L E OF GLUTAMATE UPTAKE IN SYNAPTIC TRANSMISSION 6.1. OVERVIEW In order to understand the transmission at glutamatergic synapses, which represent the majority of the excitatory synapses in the CNS (Ottersen and Storm-Mathisen, 1984), it is necessary to understand how the concentration of glutamate changes after synaptic release. From the moment glutamate is released, it will diffuse from the point of release and interact with glutamate-binding proteins, which includes transporters and receptors (both of which are not only found in the synapse, but also outside it). The binding to the various proteins will reduce the concentration of free glutamate, but will also slow down the diffusion away from the site of release (see Section 6.3). The various glutamate-binding proteins have very different properties. The receptors differ with respect to affinities as well as opening and inactivation times, while the glutamate transporter subtypes differ with respect to affinities and associated ion conductances (e.g. chloride-channel activity). The glutamate concentrations achieved at various locations from the release site as well as how quickly the concentrations change, will determine where glutamate receptors and transporters are activated as well as which subtypes and how many receptors are activated. Further, these parameters will also determine if, or to what extent, release of glutamate at one synapse leads to the activation of receptors at neighboring synapses (so-called intersynaptic crosstalk). Although significant progress has been made in recent years, essential pieces of information are still lacking. This includes information on the amount of glutamate released (see below) and on the densities of EAAC and the nerve terminal glutamate transporter. 6.2. THE TIME COURSE OF GLUTAMATE IN THE SYNAPTIC CLEFT Because mathematical models (e.g. Holmes, 1995; Clements, 1996; Kleinle et al., 1996; Barbour and H~iusser, 1997) suggest that passive diffusion alone causes a rapid decline in the glutamate concentration in the synaptic cleft after release and because the glutamate transporters have a long cycling time (12-70 ms: Wadiche et al., 1995b; Kavanaugh, 1999), 243

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it has been argued (e.g. Otis et al., 1996) that glutamate uptake is only important for the slow components of glutamate removal and for the ambient glutamate levels. However, if a very high number of glutamate transporters are present close to the release sites, they could immobilize free glutamate on a submillisecond time scale by binding rather than by transport (Tong and Jahr, 1994). The notion that glutamate quickly reaches the synaptic perimeter is supported by the detection of astroglial glutamate transporter associated anion currents in less than 1 ms after release of glutamate (Bergles et al., 1997; Bergles and Jahr, 1997). Similarly, glutamate transporter associated currents are detected in cerebellar Purkinje cells 0.1 ms after release from climbing fibers suggesting that glutamate transporters are present very near release sites (Otis et al., 1997; see EAAT4 in Fig. 1). Inhibition of glutamate uptake prolongs the EPSC at some synapses (Barbour et al., 1994; Takahashi et al., 1996) and also leads to reduced release (Maki et al., 1994) due to increased activation of presynaptic metabotropic receptors inhibiting glutamate release. Modeling of glutamate diffusion and the role of transporters is still difficult because the estimates for the peak concentration of glutamate in the synaptic cleft varies from 0.014 to 11 mM (Harris and Sultan, 1995; Schikorski and Stevens, 1997). Accordingly, there is no consensus with regard to the receptor occupancy. 6.3. DENSITIES OF GLUTAMATE TRANSPORTERS AND PARADOXICAL EFFECTS Recent evidence suggests that the glutamate-binding capacity of the known glutamate transporters (15,000 and 23,000 transporter molecules per ~m 3 in the stratum radiatum of hippocampus CA1 and the molecular layer of cerebellum, respectively) is significant compared to the release capacity (Lehre and Danbolt, 1998). The average densities of glutamatergic synapses in the stratum radiatum of hippocampus CA1 and the cerebellar molecular layer are 0.9-1.3 ~m -3 (Woolley and McEwen, 1992) and 0.8 ~m -3 (Harvey and Napper, 1991), respectively. One synaptic vesicle is believed to contain 400-5000 molecules (Clements, 1996; Barbour and Hfiusser, 1997; Schikorski and Stevens, 1997). The average sustainable release capacity has been estimated to 2 vesicles s -1 (Stevens and Tsujimoto, 1995: each average central synapse has about 20 release sites which each need about 10 s to refill). Kinetic simulations of glutamate diffusion (Rusakov and Kullmann, 1998) predict that high densities of transporters with long cycling times can lead to paradoxical effects. Binding by the transporters may rapidly reduce the extrasynaptic concentration of free glutamate after the first millisecond, but in binding glutamate, the transporters also slow down its diffusion away from the site of release. This is important because the binding is reversible. Thus, a high density of transporters on e.g. a glial process apposed to or ensheathing a synapse may trap glutamate escaping from the cleft and give it a chance to re-enter the cleft upon unbinding from the transporters. 6.4. INTERSYNAPTIC CROSSTALK Another concept which complicates the interpretation of the roles of the transporters, is the idea of intersynaptic crosstalk. It has been suggested (Kullmann and Asztely, 1998) that the reason why larger quantal contents are sensed by NMDA receptors than by AMPA receptors is that glutamate is spilling over from one synapse to another. At the neighboring synapse, the concentration of glutamate is low. Because NMDA receptors have higher affinities than AMPA receptors, it follows that NMDA receptors will more readily be activated than the 244

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AMPA receptors. This difference in quantal content is temperature-sensitive, being reduced at 37~ as compared to room temperature (Asztely et al., 1997). The authors suggest that the temperature effect could be due to glutamate transporters' more efficiently preventing spillover at 37~ This interpretation is strengthened by the finding that the temperature effect is sensitive to dihydrokainic acid, a glutamate uptake blocker. To understand the role of glutamate transporters in limiting intersynaptic crosstalk, it is necessary to find out where they are located in relation to the release sites. Since most of the transporters are on astrocytes (GLAST and GLT as well as some of the EAAC) or on neuronal membranes facing astrocytes (EAAT4) (Dehnes et al., 1998; Lehre and Danbolt, 1998), the question of whether the transporters contribute significantly to preventing glutamate from reaching neighboring synapses is more or less the same as asking where the astrocytic processes are in relation to the release sites and the diffusion barriers (unless the nerve terminal glutamate transporter or novel postsynaptic transporters, as explained above [see Section 2], contribute significantly). In the molecular layer of the cerebellum, glutamatergic synapses are often almost completely ensheathed by glia, and neighboring synapses are thereby usually separated by astrocytic processes (expressing high densities of GLAST and GLT). In contrast, most of the synapses in hippocampus are contacted by an astrocytic process which usually covers less than half of the synaptic circumference (Spacek, 1985). Further, only 33% of neighboring synapses have an astrocytic process between them (Ventura and Harris, 1999). This implies (Lehre and Danbolt, 1998) that in the cerebellum, glutamate transporters are usually in the position to interact with glutamate diffusing out of a typical synapse (i.e. parallel fiber synapse on Purkinje cell spines) before it enters the cleft of the neighboring synapse. This may not be the case at typical hippocampal synapses (i.e. Schaffer collateral synapses on pyramidal cell spines). Before jumping to conclusions, one should keep in mind that all the structures in the tissues are dynamic. Both dendritic spines (Fifkova, 1985; Fischer et al., 1998) and astrocytic processes (Wenzel et al., 1991) are able to change their forms by contraction and distention. Recent studies suggest that astrocytes preferentially extend their processes to the active synapses (Ventura and Harris, 1999) and that the dimensions of dendritic spines are regulated, in part, by glutamate receptors (Korkotian and Segal, 1999). Further, glutamate transporter (see Section 5) and receptor (Rao and Craig, 1997) densities are subject to various kinds of regulation. The receptor expression is modulated by activity (e.g. Lissin et al., 1998; Fava et al., 1999; Quinlan et al., 1999), by steroids (Gibbs et al., 1999) and other factors. The modulatory mechanisms include alternative splicing of mRNA and trafficking of the proteins to the cell surface and the cytoplasm (e.g. Lomeli et al., 1994; Zhao et al., 1998; Okabe et al., 1999; Roche et al., 1999). One should also keep the possible paradoxical effects of glutamate binding to transporters in mind (see Section 6.3).

7. CONCLUDING REMARKS

For a long time glutamate uptake was regarded as a simple drainage system which is important for securing glutamatergic neurotransmission, but which does not take active part in the signal transduction itself. This picture has now changed. Although we do not yet know exactly how glutamate transporters are involved in the process, it is becoming clear that they play refined roles. More information on the glutamate uptake system is necessary if glutamatergic neurotransmission shall be properly understood. Thus, this has not only theoretical importance, but is likely to be important for understanding a variety of diseases. 245

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8. ABBREVIATIONS

EAAC EAAT GABA GLAST GAT1 GLT GLYT NMDA SDS SDS-PAGE

rabbit glutamate transporter (Kanai and Hediger, 1992) excitatory amino acid transporter (synonym to glutamate transporter) y-aminobutyric acid rat glutamate transporter (Storck et al., 1992) GABA transporter 1 (Guastella et al., 1990) rat glutamate transporter (Pines et al., 1992) glycine transporter N-methyl-D-aspartate sodium dodecyl sulfate sodium dodecyl sulfate-polyacrylamide gel electrophoresis

9. ACKNOWLEDGEMENTS

This work was supported by the Norwegian Research Council. I would like to thank Jon Storm-Mathisen for critical reading of the manuscript.

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Glutamate neurotransmission in the m a m m a l i a n inner ear S. USAMI, A. MATSUBARA, S. FUJITA, Y. TAKUMI AND O.E OTTERSEN

1. I N T R O D U C T I O N Previous chapters in this volume have dealt with central glutamate synapses and the principles underlying their structural and molecular organization. An important question is whether these principles are applicable also to peripheral glutamate synapses, which engage cell types that differ from central neurons in terms of morphology and embryological origin. The present chapter aims at addressing this issue by focusing on a set of putative glutamatergic synapses in the inner ear. The inner ear contains the cochlea and vestibular endorgans, the sensory organs for hearing and equilibrium. The receptor cells in these organs are named hair cells since they are equipped with stereocilia that respond to mechanical stimulation. The first synapse in the sensory pathways is that between hair cells and primary afferent neurons. Glutamate has long been considered as the most likely neurotransmitter candidate in this synapse. This view is supported by a series of pharmacological and electrophysiological studies (reviewed by: Bobbin, 1979; Bledsoe et al., 1988; Ehrenberger and Felix, 1991; Puel et al., 1991; Puel, 1995; Ottersen et al., 1998) and by recent neurochemical and immunocytochemical investigations that will be discussed here. The picture that has emerged from the latter investigations is that the afferent hair cell synapse has several features in common with central glutamate synapses. Many of the proteins known to be involved in signal transduction and transmitter metabolism at most central synapses also occur in the afferent hair cell synapse, and with an analogous compartmentation. However, important differences also exist, notably in regard to the molecular mechanisms underlying transmitter release.

2. GLUTAMATE IN HAIR CELLS One of the criteria to be fulfilled by a neurotransmitter is that it must be present in the presynaptic element. However, a presynaptic localization is no proof of transmitter identity. Glutamate, in particular, serves multiple functions in cell metabolism and may be quite abundant even in cells that do not use glutamate as a neurotransmitter (see Chapter 1 for a thorough discussion of this issue). Biochemical analyses of micro-dissected samples from the hair cell region in the organ of Corti revealed significant amounts of glutamate (Godfrey et al., 1976, 1986), indicating that Handbook of Chemical Neuroanatomy, Vol. 18: Glutamate O.E Ottersen and J. Storm-Mathisen, editors (g) 2000 Elsevier Science B.V. All rights reserved.

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this amino acid might be enriched in hair cells. More direct evidence of this was obtained in light microscopic studies which demonstrated glutamate immunoreactivity in cochlear as well as vestibular hair cells (Altschuler et al., 1989; Dem~mes et al., 1990; Usami et al., 1992). Detailed analyses at the light and electron microscopic levels have displayed a highly differentiated pattern of glutamate-like immunoreactivity in the organ of Corti (Fig. 1). Although supporting cells are generally less strongly immunoreactive than the hair cells, their staining intensity varies over a relatively wide range. Inner pillar cells, inner phalangeal cells,

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and border cells are moderately immunoreactive, while supporting cells further r e m o v e d from the hair cell region (the Hensen cells and inner sulcus cells) are almost devoid of labelling (Fig. 1). However, the glutamate staining intensity does not follow a strict proximo-distal gradient since the Boettcher cells are clearly more strongly labelled than the adjacent Hensen cells (Fig. 1A,C). The sub-basilar tympanic cells that line the basilar m e m b r a n e are atypical in that they show a strong glutamate immunoreactivity that even exceeds that of the hair cells (Fig. 1A,C). The functional significance of this finding is not clear but it underscores the fact that a high glutamate level cannot always be equated with a transmitter pool (see Chapter 1). The hair cells were also less intensely labelled than the terminals of the putatively glutamatergic parallel fibres of the cerebellum (Fig. 1C). This difference m a y be due to methodological factors: although cerebellar and cochlear sections were obtained from the same animals and incubated simultaneously, the cochlear cells m a y have suffered a greater loss of free amino acids because of less favourable fixation conditions (Usami et al., 1992). It must also be pointed out that the strength of the glutamate i m m u n o g o l d signal in central synapses is positively correlated to the density of synaptic vesicles (Ji et al., 1991; also see Chapter 1). Since the hair cells contain few vesicles per unit volume c o m p a r e d to parallel fibre terminals one would expect the cytoplasmic glutamate concentration to be lower. However, a word of caution is required at this point. Although i m m u n o g o l d particles signalling glutamate are associated with the synaptic vesicles near the base of the hair cells (Matsubara et al., 1996), conclusive evidence for an enrichment of glutamate in these vesicles is still pending. Formally one must leave open the possibility that hair cell transmission is m e d i a t e d by a glutamate receptor agonist different from glutamate. Mitochondria of hair cells were strongly i m m u n o l a b e l l e d for glutamate (Fig. 1B). This is consistent with the idea that these organelles are responsible for the synthesis of transmitter glutamate from glutamine (Chapter 1; also see below).

Fig. 1. Presynaptic localization of glutamate in hair cells. (A) Light micrograph showing the distribution of glutamate immunoreactivity in the organ of Corti (guinea pig). Inner (IHC) and outer (OHC) hair cells display strong immunoreactivity, as do the sub-basilar tympanic cells (SBT). Supporting cells are generally less intensely immunoreactive than the hair cells, and particularly weak labelling occurs in inner sulcus cells (ISC) and Hensen cells (HC). Other abbreviations: TM = tectorial membrane; BD = border cells; IPH = inner phalangeal cells; IPC and OPC = inner and outer pillar cells; DC -- Deiters cells; BC -- Boettcher cells. Frame shows area enlarged in B. Inset in A: cross-section of 'sandwich' containing 8 brain sections (dark lanes) alternating with 7 sections of resin-embedded test conjugates (aldehyde conjugates of brain protein and GABA [1], glutamate [2], taurine [3], glycine [4], no amino acid [5], aspartate [6], and glutamine [7]). This cross-section was incubated together with the section of the organ of Corti and shows that the antibody stains the glutamate conjugates exclusively (lane 2). (B) Electron micrograph of ultrathin section from the area indicated in A. Postembedding glutamate immunolabelling produced a high density of gold particles in the inner hair cells (IHC), and progressively lower labelling intensities in the border cells (BD) and inner sulcus cells (asterisk). (C) Quantitative analysis of preparations similar to that in B. S.E.M. and number of observations are indicated for each column. The sections of the organ of Corti were incubated together with sections of the cerebellum (obtained from the same animal), thus allowing comparison with the gold particle density in parallel fibre terminals (cer. par. f.) and astrocyte processes (cer. glial c.). Middle panel: test section similar to that in A (inset) but ultrathin and immunogold labelled. Standard abbreviations for amino acids. Note that selectivity of the glutamate antibody is maintained at the EM level (small particles). This particular test section was also labelled for glutamine (large particles) using a sequential double labelling procedure. Scale bar in B is 0.5 Ixm.

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3. A GLUTAMATE-GLUTAMINE CYCLE IN THE INNER EAR? GLUTAMINE SYNTHETASE AND GLUTAMATE TRANSPORTERS

Chapter 1 discussed the large body of data that points to the existence of a glutamateglutamine cycle in the CNS. This cycle is driven by the two enzymes phosphate-activated glutaminase (PAG) and glutamine synthetase (GS). The former enzyme converts glutamine to glutamate while the latter enzyme catalyses the opposite reaction, from glutamate to glutamine. The two enzymes are concentrated in neurons and glia, respectively, and would therefore cause a cycling of carbon skeletons between these cellular compartments if mechanisms for cellular exchange of glutamate and glutamine were available. Ample experimental evidence indicates that such an exchange indeed occurs although it is now recognized that the cycle is far from stoichiometrically perfect (see Chapter 1). Molecules that mediate uptake of glutamate in glial cells have been identified and cloned (see Chapter 8 by Danbolt, this volume). The glutamate transporters GLAST and GLT1 (EAAT1 and EAAT2 according to new nomenclature) are both expressed by astrocyte plasma membranes although their relative concentration varies among brain regions. These transporters are believed to help remove synaptically released glutamate from the extracellular space and form an integral part of the glutamate-glutamine cycle. The molecular mechanisms responsible for the transfer of glutamine from astrocytes to neurons have been obscure until the recent discovery and cloning of two glutamine carriers that mediate efflux of glutamine from astrocytes (Broer et al., 1999; Chaudhry et al., 1999). Carriers for glutamine uptake have been characterized (see Tamarappoo et al., 1997, and references therein) but not yet cloned. Is there any evidence of a glutamate-glutamine cycle in the inner ear? One important piece of evidence came with the demonstration that the level of glutamine immunoreactivity is lower in hair cells than in the adjoining supporting cells (Fig. 2; Usami and Ottersen, 1995). Since glutamate immunoreactivity shows the complementary distribution (Figs. 1 and 2) the glutamate/glutamine ratio must be much higher in hair cells than in supporting cells. Such differences in the glutamate/glutamine ratio have also been demonstrated between neurons and astrocytes in the CNS (Ottersen et al., 1992) and are considered a hallmark of glutamate-glutamine cycling. If supporting cells are engaged in a glutamate-glutamine cycle analogous to that in the CNS one would expect that they contain glutamine synthetase and at least one glutamate transporter. Glutamine synthetase has been demonstrated in vestibular supporting cells (Takumi et al., 1997) and in the same study it was shown by double immunofluorescence that these cells also express the glutamate transporter EAAT1. The hair cells were negative for either antigen. Immunogold labelling with antibodies to EAAT1 confirmed this result and revealed gold particles along supporting cell plasma membranes in the vestibular epithelium (Fig. 2E) as well as in the organ of Corti (Fig. 3B; also see Furness and Lehre, 1997). In both sensory organs the concentration of EAAT1 molecules is higher in those plasma membrane domains that face the synaptic region than in membrane domains more distant to the synaptic sites (Figs. 2E and 3B; also see Takumi et al., 1997). This is analogous to observations in the CNS (Chaudhry et al., 1995) and suggests that glutamate transporters are expressed according to demand. Hence, mechanisms must exist that serve to anchor EAAT1 at specific membrane domains of individual supporting cells. EAAT1 has been found in supporting cells apposed to the inner hair cells in rat and guinea pig (Furness and Lehre, 1997) as well as mouse (Hakuba et al., 2000). In contrast, no significant EAAT1 immunolabelling occurs in supporting cells in the vicinity of the outer hair cells. This agrees with the idea that EAAT1 is expressed according to demand, since 258

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Fig. 2. Amino acid compartmentation and putative glutamate-glutamine cycle in the vestibular epithelium. Immunogold particles signalling glutamate (A) are enriched in hair cells (HC) relative to supporting cells (SC), whereas the converse is true for particles signalling glutamine (C). This pattern could be confirmed by quantitative analysis (D, F) and is compatible with a differential distribution of the respective synthesizing enzymes (phosphate-activated glutaminase [PAG] in hair cells and glutamine synthetase [Gln-synthetase]in supporting cells). The two metabolic pathways may be coupled through a glutamate-glutamine cycle (B), an assumption that is supported by the finding of EAAT1 (GLAST) immunoreactivity (arrows) in supporting cell membranes (E). (This cycle is not stoichiometrically perfect; see text and Chapter 1.) Asterisks in A, C and E indicate nerve chalices. Note in D and F that the two types of hair cell (HCI and HCII) show comparable labelling intensities (expressed as number of gold particles/Ixm2). S.E.M. and number of observations are indicated. Asterisks in D and F denote values significantly different from values for supporting cells (P < 0.0.1, Student's t-test). Scale bars: 1 Ixm in A and C, 0.5 Ixm in E.

there is evidence that outer hair cells are incapable of glutamate release under physiological conditions (for references see Matsubara et al., 1996). The results discussed so far indicate that glutamate released at the afferent synapses of vestibular and cochlear hair cells may be taken up by adjacent supporting cells through the glutamate transporter EAAT1. The supporting cells may then convert glutamate into glutamine by glutamine synthetase (although this enzyme has yet to be demonstrated in cochlear supporting cells; see Eybalin et al., 1996). Taken together, the available data on glutamate compartmentation, metabolism, and transport lend support to the hypothesis that the inner ear is endowed with a glutamate-glutamine cycle similar to that assumed to operate in the CNS. However, to 259

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Fig. 3. Phosphate-activated glutaminase (PAG) and glutamate transporter (EAAT1/GLAST) in the organ of Corti. (A) Mitochondria of inner hair cells (IHC) exhibit strong immunogold labelling for PAG, whereas mitochondria (asterisk) in adjacent supporting cells (SC) contain few or no gold particles. (B) GLAST immunoreactivity (small arrows) is concentrated in supporting cell plasma membranes surrounding the synaptic areas at the base of the inner hair cells. Other abbreviations: aft -- afferent nerve fibre; eft - efferent nerve fibre. Scale bars: 0.5 I~m in A, 1 I~m in B.

complete the cycle the hair cells must contain a glutaminase activity, allowing them to form glutamate from glutamine. Whether this is the case will be discussed in the next paragraph.

4. DISTRIBUTION OF PHOSPHATE-ACTIVATED GLUTAMINASE IN THE INNER EAR

Phosphate-activated glutaminase (PAG) is assumed to be responsible for most of the glutaminase activity in the CNS (Kvamme, 1984). This enzyme has been found in a large number of neuronal pathways, some of which are thought to be glutamatergic on other grounds (see Chapter 7 by Kaneko of this volume). By use of postembedding immunogold cytochemistry it was recently shown that PAG is virtually restricted to mitochondria (Laake et al., 1999). However, the mitochondrial labelling intensity varied over a wide range, depending on the identity of the cell compartment. Cerebellar mossy fibre terminals contained strongly immunoreactive mitochondria, whereas other putative glutamatergic fibres (such as the parallel fibres) contained mitochondria with substantially lower particle densities (Laake et al., 1999). This suggests that the mechanisms for transmitter replenishment may be less uniform than previously, assumed and that some glutamatergic terminals may depend heavily on other sources of glutamate than PAG activity. Glial cell mitochondria were devoid of specific PAG 260

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Fig. 4. Phosphate-activated glutaminase (PAG) in the vestibular epithelium. Strong immunogold labelling for PAG occurs in the mitochondria of type I hair cell (HC), whereas mitochondria of supporting cells (SC, arrowheads) are virtually unlabelled. Asterisks indicate nerve chalice. Scale bar: 0.5 t*m. Modified from Takumi et al. (1999a).

labelling, in agreement with the classical concept of the glutamate-glutamine cycle. Using the same antibody as Laake et al. (1999), Takumi et al. (1999a) demonstrated a selective enrichment of PAG in cochlear and vestibular hair cell mitochondria (Fig. 3A, Fig. 4). The gold particle density over supporting cell mitochondria was less than 15% of that in hair cells. This underscores the similarity between the CNS and inner ear as regards metabolic compartmentation. Triple immunogold labelling for glutamate, glutamine, and PAG revealed a positive correlation between the glutamate/glutamine ratio and the level of PAG immunoreactivity (Takumi et al., 1999a). This indicates that the PAG antibodies identify a functional pool of this enzyme. Using small gold particles for optimum resolution it could be shown that PAG is likely to be associated with the inner mitochondrial membrane, although there may be an additional enzyme pool in the mitochondrial matrix. The outer mitochondrial membrane was invariably unlabelled. No differences were found between basal and apical parts of the hair cells with respect to the PAG immunolabelling intensity of their mitochondria (Takumi et al., 1999a). This suggests that the hair cells are unable to maintain an intracellular gradient of PAG and that the synaptic pole of these cells is not defined at any step prior to vesicular transmitter uptake. 261

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5. G L U T A M A T E R E L E A S E The presynaptic specialization of the hair cells is characterized by an electron dense body (Friedmann and Ballantyne, 1984). This is localized near the centre of the synapse and is surrounded by synaptic vesicles. As discussed below, it is believed that synaptic release occurs mainly or exclusively at the site of the presynaptic dense body (Furukawa et al., 1982; Parsons et al., 1994). The morphological dissimilarity between the hair cell synapse and most central synapses indicates different release mechanisms. In agreement, synaptotagmin I and II, which are considered to play major roles in neurotransmitter release at central synapses, appear to be absent from inner and outer hair cells (Safieddine and Wenthold, 1999). The same is true of synapsin and synaptophysin. However, the hair cells do contain syntaxin 1, the synaptic membrane-associated protein SNAP-25, and the vesicle-associated membrane protein VAMP (Safieddine and Wenthold, 1999). The hair cells also differ from most central synapses in regard to their complement of voltage-dependent Ca2+-channels. In the hair cells, transmitter release is triggered by the opening of dihydropyridine-sensitive L-type channels, although these have properties that set them apart from L-type channels in other cells (Hudspeth and Lewis, 1988; Roberts et al., 1990). A detailed discussion of this issue can be found elsewhere (Ottersen et al., 1998).

6. G L U T A M A T E R E C E P T O R S A large variety of glutamate receptors of every major family has been demonstrated at the protein or mRNA level in spiral and vestibular ganglion cells (Kuriyama et al., 1993, 1994; Fujita et al., 1994; Demames et al., 1995; Niedzielski and Wenthold, 1995; also see Fig. 6A). The challenge has been to identify the receptor types that are expressed at the afferent synapse itself and which take part in hair cell transmission. 6.1. AMPA RECEPTORS To date the only glutamate receptors that have been consistently localized to the postsynaptic specialization of the afferent hair cell synapses are the AMPA receptors GluR2/3 and 4 (Matsubara et al., 1996, 1999). Using immunogold techniques, these subunits were found postsynaptic to three types of hair cell (Figs. 5 and 6): the inner hair cells in the organ of Corti (Matsubara et al., 1996) and type I and type II hair cells in the vestibular epithelium (Matsubara et al., 1999). No receptor immunolabelling could be detected postsynaptic to the outer hair cells in the cochlea (Fig. 5C). Although negative observations should be interpreted

Fig. 5. Distribution of AMPA receptors in the organ of Corti. (A, B) Immunogold particles in afferent hair cell

synapses after incubation with an antibody recognizing GluR2 and GluR3. The particles are concentrated in the postsynaptic specialization (between arrowheads). The synapse in B has been cut through the presynaptic dense body (asterisk); that in A has been cut off centre. Both are from inner hair cells (IHC). Although not evident in these individual profiles, a statistical analysis of a large sample of synapses revealed a higher gold particle density near the margin of the synapse than more centrally (see text). Arrows indicate synaptic vesicles. (C) Region at the base of the outer hair cells (OHC). This section was processed together with those represented in A and B, but does not exhibit any GluR2/3 immunoreactivity. Other abbreviations: aft = afferent nerve fibre; eft = efferent nerve fibre, m = mitochondrion. 262

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Q.,;I,~i::;!ID Fig. 6. Distribution of AMPA receptors in the vestibular endorgan. (A) Light micrograph showing intense GluR2/3 immunoreactivity in the vestibular ganglion cells (peroxidase-antiperoxidase procedure). (B-D) Electron micrographs showing GluR2/3 immunogold labelling in synapses between hair cell type I (HCI) and nerve chalice (B, C), between hair cell type II (HCII) and afferent nerve fibre (D, left), and between HCII and the outer face of the nerve chalice (D, right). Small arrows indicate gold particles; asterisks denote nerve chalices, aft = afferent nerve fibre. Scale bars: 50 Ixm in A, 0.2 Ixm in B-D.

with due caution (Nusser et al., 1998; Takumi et al., 1999b), it should be pointed out that there is still no evidence that acoustic stimulation elicits a response in the afferent fibres that lead to the outer hair cells (Patuzzi and Robertson, 1988). The functional silence may reflect the absence of appropriate postsynaptic receptors. In addition or alternatively, the outer hair cells may be incapable of glutamate release within the normal range of stimulus intensities. This is because their voltage-dependent CaZ+-channels seem to have a threshold that is beyond the maximum depolarization of the outer hair cell membrane (compare Patuzzi and Robertson, 264

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1988, with Nakagawa et al., 1991a). In agreement, the outer hair cells have been found to exhibit a low rate of vesicle recycling compared to the inner hair cells (Siegel and Brownell, 1986). A detailed analysis has been performed of the AMPA receptor distribution at the afferent synapse of inner hair cells (Matsubara et al., 1996). Using quantitative postembedding immunocytochemistry and serial sections (to identify the centre of the synapse) it was found that the concentration of GluR2/3 was higher near the periphery of the postsynaptic density than more centrally. The same gradient was identified for the GluR4 subunits which appeared to be colocalized with GluR2/3 throughout the synapse. The functional implication of this arrangement is unclear. However, it should be noted that in the afferent hair cell synapse, exocytosis is thought to occur mainly or exclusively at the site of the presynaptic dense body (Furukawa et al., 1982; Parsons et al., 1994). This implies that glutamate has to diffuse more than 200 nm in the lateral direction to reach the most peripheral receptors of this large synapse (Matsubara et al., 1996). The enrichment of receptors at the periphery might help compensate for the lateral attenuation of glutamate and ensure an even density of open receptor channels throughout the postsynaptic specialization. An analysis of the gold particle distribution along an axis perpendicular to the postsynaptic specialization revealed a distinct peak over the postsynaptic density (Matsubara et al., 1996). However, for GluR4 but not GluR2/3, an additional but smaller peak was observed over the presynaptic membrane. This suggests that the inner hair cells are endowed with a small pool of AMPA receptors containing GluR4 subunits. Physiological evidence of presynaptic AMPA receptors has been obtained for type I vestibular hair cells (Devau et al., 1993) and it could well be that they act as autoreceptors. This would differ from the situation in the CNS where autoreceptors are typically insensitive to AMPA (Chittajallu et al., 1996). In the vestibular epithelium, AMPA receptors (GluR2/3 and GluR4) were found in three types of synapse (Matsubara et al., 1999): between type I cells and nerve chalices (Fig. 6B,C), between type II cells and afferent fibres (Fig. 6D), and between type II cells and the outer face of nerve chalices (Fig. 6D). Some nerve chalices are thus likely to receive a glutamatergic input from type I as well as type II hair cells. These chalices may correspond to the dimorphic units characterized by Goldberg et al. (1990). As the individual synaptic contacts between type I cells and nerve chalices are very small they rarely exhibited more than two or three gold particles (Fig. 6B,C). The two other types of contact are more extensive and hence the number of particles per contact was larger (Fig. 6D). Immunoreactivity for GluR1 was not observed in the afferent hair cell synapses in the vestibular epithelium, nor at the inner or outer hair cell synapses in the organ of Corti (Matsubara et al., 1996, 1999). The immunoincubations on which this observation was based produced strong labelling of hippocampal synapses, indicating that the lack of labelling was not merely due to a methodological artefact. In agreement, while ganglion cells have been shown to contain GluR2-4 and their respective mRNAs (Ryan et al., 1991; Safieddine and Eybalin, 1992; Kuriyama et al., 1994; Luo et al., 1995; Niedzielski and Wenthold, 1995; Usami et al., 1995), there is no evidence in the literature that adult ganglion cells express significant amounts of GluR1. This pattern may be phylogenetically conserved since auditory ganglion cells in the pigeon exhibit immunoreactivity for GluR2/3 and GluR4 but not for GluR1 (Reng et al., 1999). We can thus conclude from the immunoelectron microscopical data that AMPA receptors composed of GluR2/3 and GluR4 subunits are likely to be involved in afferent hair cell transmission. This would be in line with physiological studies. Patch-clamp analyses have identified functional AMPA receptors in isolated spiral ganglion cells (Nakagawa et al., 265

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1991b; Ruel et al., 1999), and antisense knockdown of GluR2 was shown to reduce the compound action potential and diminish spontaneous activity of single auditory nerve fibres (D'Aldin et al., 1998). The effect of the antisense probe was confirmed by demonstrating a reduction in GluR2/3 immunoreactivity in ganglion cells. 6.2. OTHER TYPES OF GLUTAMATE RECEPTOR NMDA receptors and NMDA receptor mRNAs have been demonstrated in the cell bodies of spiral ganglion cells (Kuriyama et al., 1993; Fujita et al., 1994; Niedzielski and Wenthold, 1995). It is still unclear, however, whether functional NMDA receptors are expressed at the afferent hair cell synapse. Matsubara et al. (1996) found no evidence of such expression under experimental conditions that produced intense immunogold labelling of non-NMDA receptor isoforms (see above). Pharmacological experiments have provided conflicting data but recent electrophysiological studies seem to argue against a role for NMDA receptors in the afferent transmission of cochlear hair cells (see discussion in Ottersen et al., 1998). Notably, no NMDA response could be obtained in isolated spiral ganglion cells of guinea pigs (Nakagawa et al., 1991b; Ruel et al., 1999). This does not rule out that NMDA receptors may be expressed at specific stages during development (Knipper et al., 1997) or in pathological conditions (Puel et al., 1997; D'Aldin et al., 1997). In fact, glutamate has been proposed to play a neurotrophic role and to promote repair processes through activation of NMDA receptors (Puel et al., 1997). In the vestibular epithelium, the NMDA receptor subunit NR-1 has been immunolocalized to the afferent chalices of type I hair cells (Ishiyama et al., 1999). No labelling was found in the boutons innervating type II hair cells. The latter study was performed in the chinchilla. Spiral ganglion cells have been shown to express several metabotropic glutamate receptor isoforms and their respective mRNAs (Safieddine and Eybalin, 1995; Niedzielski et al., 1997). Kleinlogel et al. (1999), working in the guinea pig, reported a long-lasting increase in afferent firing after application of an mGluR1 agonist and concluded that mGluR1 could be involved in peripheral auditory processing. Evidence from the frog vestibular endorgan suggests that mGluR1 is expressed in hair cells and that it could act as an autoreceptor (Guth et al., 1998). In support of this view, Guth et al. (1998) observed that the mGluR1 agonist 1-aminocyclopentane-trans-l,3-dicarboxylate (ACPD) failed to increase afferent firing under experimental conditions known to inhibit transmitter release. Hair cells have also been shown to express high levels of the deltal glutamate receptor (Safieddine and Wenthold, 1997). This receptor was restricted to the inner hair cells in the organ of Corti but occurred in both types of hair cell in the vestibular epithelium. Evidence was also obtained of deltal expression in spiral as well as vestibular ganglion cells (Safieddine and Wenthold, 1997). The functional role of the delta l receptor is still unclear.

7. PATHOLOGY OF THE GLUTAMATERGIC SYNAPSE A major cause of hearing loss is acoustic trauma. Experimental exposure to loud noise causes mechanical damage to the outer hair cells and swelling of the afferent fibres below the inner hair cells (Robertson, 1983; Saunders et al., 1985). The morphological changes of the afferent fibres resemble those observed after application of glutamate agonists (Puel et al., 1991; Puel, 1995). Since noise is known to cause an efflux of excitatory amino acids from the cochlea (for references, see Jager et al., 1998) it has been hypothesized that excitotoxic mechanisms 266

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might contribute to noise-induced hearing loss. This would be in line with the protective effect of glutamate receptor antagonists (Puel et al., 1998). The above hypothesis was recently tested in mice with a knockout of EAAT1, the major glutamate transporter in the organ of Corti (Furness and Lehre, 1997; Takumi et al., 1997). Compared to the wild type mice, mice deficient in EAAT1 showed a more pronounced extracellular accumulation of glutamate after sound exposure, and a more severe hearing loss (Hakuba et al., 2000). This finding supports the idea that the extracellular glutamate level may rise to ototoxic levels under certain pathological conditions. It also attests to the importance of EAAT 1 in inner ear function. As in the CNS, glutamate transporters may not only help terminate synaptic transmission but may also act as a safeguard against a harmful buildup of extracellular glutamate. It serves to illustrate this point that kanamycin m an ototoxic drug - - has been found to increase the expression of EAAT1 in the inner ear (Matsuda et al., 1999). This change is likely to be compensatory and to have a neuroprotective effect. A possibility to be tested is whether EAAT1 expression is induced by high glutamate concentrations, such as those that are likely to occur when glutamate is lost from collapsing hair cells (Matsuda et al., 2000). It has been suggested that glutamate neurotoxicity is involved in a wide range of pathological states in addition to those discussed above (Pujol et al., 1993; Puel, 1995; Basile et al., 1996). Examples are neural presbyacusis, some forms of peripheral tinnitus, and ischemia. As to ischemia, this condition has been shown to be associated with an increased glutamate concentration in the perilymph (Hakuba et al., 1997; Haruta et al., 1998) which may be secondary to an efflux of glutamate from hair cells and supporting cells (Matsubara et al., 1998). The ischemia-induced swelling of afferent fibres can be prevented by glutamate receptor antagonists (Pujol et al., 1993). The molecular mechanisms underlying glutamate toxicity in the inner ear are not known but an increased NO production (Sunami et al., 1999a) or impaired cystine-glutamate exchange (Sunami et al., 1999b) may be involved. The importance of excitotoxicity as a pathogenetic factor in inner ear disease calls for the development of specific neuroprotective drugs. Piribedil, a D2 dopamine receptor agonist, was reported to counteract radial dendritic swelling following transient ischemia (Pujol et al., 1993; D'Aldin et al., 1995). This points to bromocriptine as an interesting prototype in the development of otoprotective drugs: bromocriptine is not only a D2 dopamine receptor agonist but has also been shown to stimulate glutamate transport through a dopamine-receptor-independent mechanism (Yamashita et al., 1995; Yamashita et al., 1998). Interestingly, dopamine alone has little effect on the spontaneous firing rate of afferent fibres from inner hair cells but significantly depresses firing induced by coapplication of NMDA or AMPA (Oestreicher et al., 1997). Dopamine is one of several neuroactive substances that have been identified in the efferent olivocochlear fibre system (Jones et al., 1987; Usami et al., 1988; Eybalin et al., 1993; D'Aldin et al., 1995; Gil-Loyzaga, 1995; Gaborjan et al., 1999).

8. C O N C L U S I O N The afferent hair cell synapse now emerges as one of the best characterized glutamate synapses in mammals. Its attractiveness as a model synapse derives from the fact that it is well defined morphologically and functionally and can be studied in relative isolation from other synapses. Experimental studies of the hair cell synapse should thus allow us to investigate the physiological and pathophysiological role of individual synaptic proteins, a possibility that is now beginning to be realized by knockout and knockdown approaches. It is interesting in this regard that the afferent hair cell synapse is built according to many of the same principles 267

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as central synapses, although the cellular players are different. Notably, in the inner ear the supporting cells appear to carry out a number of tasks that depend on astrocytes in the CNS. Glutamate uptake and metabolism are two important examples. Other examples not discussed here are water and ion homeostasis, and volume regulation (Takumi et al., 1998). It is now clear that the different types of supporting cell have highly specialized functions that correlate with their structural heterogeneity and orderly arrangement in the sensory epithelium. On a general note one can conclude that glutamate neurotransmission is dependent on a functional interaction with non-neuronal cells, be it astrocytes in the CNS or supporting cells in the inner ear. The glutamate synapse is indeed a tripartite contact.

9. ACKNOWLEDGEMENTS The work reviewed in this chapter was supported by a Grant-in-Aid for Scientific Research from the Ministry of Education, Science and Culture of Japan, the Ministry of Health and Welfare of Japan, The Karoji Memorial Fund for Medical Research in Hirosaki University, Ground Research for Space Utilization promoted by NASA and Japan Space Forum, the Norwegian Research Council, Letten E Saugstad's Fund, and the Sasakawa Foundation.

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Kuriyama H, Albin RL, Altschuler RA (1993): Expression of NMDA-receptor mRNA in the rat cochlea. Hear Res 69:215-220. Kuriyama H, Jenkins O, Altschuler RA (1994): Immunocytochemical localization of AMPA selective glutamate receptor subunits in the rat cochlea. Hear Res 80:233-240. Kvamme E (1984): Enzymes of cerebral glutamine metabolism in mammalian tissues. In: Haussinger D, Sies H (Eds), Glutamine Metabolism in Mammalian Tissues. Berlin: Springer, pp 32-48. Laake JH, Takumi Y, Eidet J, Torgner IA, Roberg B, Kvamme E, Ottersen OP (1999): Postembedding immunogold labelling reveals subcellular localization and pathway-specific enrichment of phosphate activated glutaminase in rat cerebellum. Neuroscience 88:1137-1151.

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Luo L, Brumm D, Ryan AF (1995): Distribution of non-NMDA glutamate receptor mRNAs in the developing rat cochlea. J Comp Neurol 361:372-382. Matsubara A, Laake JH, Davanger S, Usami S, Ottersen OP (1996): Organization of AMPA receptor subunits at a glutamate synapse: a quantitative immunogold analysis of hair cell synapses in the rat organ of Corti. J Neurosci 16:4457-4467. Matsubara A, Kawabata Y, Takumi Y, Usami S, Shinkawa H, Haruta A, Matsuda K, Tono T (1998): Quantitative immunogold cytochemistry reveals sources of glutamate release in inner ear ischemia. Acta Otolaryngol Suppl (Stockh) 539:48-51. Matsubara A, Takumi Y, Nakagawa T, Usami S, Shinkawa H, Ottersen OP (1999): Immunoelectron microscopy of AMPA receptor subunits reveals three types of putative glutamatergic synapse in the rat vestibular end organs. Brain Res 819:58-64. Matsuda K, Ueda Y, Doi T, Tono T, Haruta A, Toyama K, Komune S (1999): Increase in glutamate-aspartate transporter (GLAST) mRNA during kanamycin-induced cochlear insult in rats. Hear Res 133:10-16. Matsuda K, Komune S, Tono T, Yamasaki M, Haruta A, Kato E (2000): A role of glutamate in drug-induced ototoxicity: in vivo microdialysis study combined with on-line enzyme fluorometric detection of glutamate in the guinea pig cochlea. Brain Res 852:492-495. Nakagawa T, Kakehata S, Akaike N, Komune S, Takasaka T, Uemura T (1991a): Calcium channel in isolated outer hair cells of guinea pig cochlea. Neurosci Lett 125:81-84. Nakagawa T, Komune S, Uemura T, Akaike N (1991b): Excitatory amino acid response in isolated spiral ganglion cells of guinea pig cochlea. J Neurophysiol 65:715-723. Niedzielski AS, Wenthold RJ (1995): Expression of AMPA, kainate, and NMDA receptor subunits in cochlear and vestibular ganglia. J Neurosci 15:2338-2353. Niedzielski AS, Safieddine S, Wenthold RJ (1997): Molecular analysis of excitatory amino acid receptor expression in the cochlea. Audiol Neurootol 2:79-91 [erratum in Audiol Neurootol 1997 Jul-Aug 2(4):231 ]. Nusser Z, Lujan R, Laube G, Roberts JD, Moln~ir E, Somogyi P (1998): Cell type and pathway dependence of synaptic AMPA receptor number and variability in the hippocampus. Neuron 21:545-559. Oestreicher E, Arnold W, Ehrenberger K, Felix D (1997): Dopamine regulates the glutamatergic inner hair cell activity in guinea pigs. Hear Res 107:46-52. Ottersen OP, Zhang N, Walberg F (1992): Metabolic compartmentation of glutamate and glutamine: morphological evidence obtained by quantitative immunocytochemistry in rat cerebellum. Neuroscience 46:519-534. Ottersen OP, Takumi Y, Matsubara A, Landsend AS, Laake JH, Usami S (1998): Molecular organization of a type of peripheral glutamate synapse: the afferent synapses of hair cells in the inner ear. Prog Neurobiol 54:127-148. Parsons TD, Lenzi D, Almers W, Roberts WM (1994): Calcium-triggered exocytosis and endocytosis in an isolated presynaptic cell: capacitance measurements in saccular hair cells. Neuron 13:875-883. Patuzzi R, Robertson D (1988): Tuning in the mammalian cochlea. Physiol Rev 68:1009-1082. Puel JL (1995): Chemical synaptic transmission in the cochlea. Prog Neurobiol 47:449-476. Puel JL, Ladrech S, Chabert R, Pujol R, Eybalin M (1991): Electrophysiological evidence for the presence of NMDA receptors in the guinea pig cochlea. Hear Res 51:255-264. Puel JL, D'Aldin C, Ruel J, Ladrech S, Pujol R (1997): Synaptic repair mechanisms responsible for functional recovery in various cochlear pathologies. Acta Otolaryngol (Stockh) 117:214-218. Puel JL, Ruel J, Gervais D'Aldin C, Pujol R (1998): Excitotoxicity and repair of cochlear synapses after noise-trauma induced hearing loss. Neuroreport 9:2109-2114. Pujol R, Puel JL, Gervais D'Aldin C, Eybalin M (1993): Pathophysiology of the glutamatergic synapses in the cochlea. Acta Otolaryngol (Stockh) 113:330-334. Reng D, Hack I, Muller M, Smolders JW (1999): AMPA-type glutamate receptor subunits are expressed in the avian cochlear hair cells and ganglion cells. Neuroreport 10:2137-2141. Roberts WM, Jacobs RA, Hudspeth AJ (1990): Colocalization of ion channels involved in frequency selectivity and synaptic transmission at presynaptic active zones of hair cells. J Neurosci 10:3664-3684. Robertson D (1983): Functional signifcance of dendritic swelling after loud sounds in the guinea pig cochlea. Hear Res 9:263-278. Ruel J, Chen C, Pujol R, Bobbin RE Puel JL (1999): AMPA-preferring glutamate receptors in cochlear physiology of adult guinea-pig. J Physiol (Lond) 518:667-680. Ryan AF, Brumm D, Kraft M (1991): Occurrence and distribution of non-NMDA glutamate receptor mRNAs in the cochlea. Neuro repo rt 2:643-646. Safieddine S, Eybalin M (1992): Co-expression of NMDA and AMPA/kainate receptor mRNAs in cochlear neurones. Neuroreport 3:1145-1148. 270

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Safieddine S, Eybalin M (1995): Expression of mGluR1 alpha mRNA receptor in rat and guinea pig cochlear neurons. Neuroreport 7:193-196. Safieddine S, Wenthold RJ (1997): The glutamate receptor subunit deltal is highly expressed in hair cells of the auditory and vestibular systems. J Neurosci 17:7523-7531. Safieddine S, Wenthold RJ (1999): SNARE complex at the ribbon synapses of cochlear hair cells: analysis of synaptic vesicle- and synaptic membrane-associated proteins. Eur J Neurosci 11:803-812. Saunders JC, Dear SP, Schneider ME (1985): The anatomical consequences of acoustic injury: a review and tutorial. J Acoust Soc Am 78:833-860. Siegel JH, Brownell WE (1986): Synaptic and Golgi membrane recycling in cochlear hair cells. J Neurocytol 15:311-328. Sunami K, Yamane H, Nakagawa T, Takayama M, Konishi K (1999a): Glutamate toxicity induced degeneration of outer hair cells with a temporal increase of nitric oxide production in the guinea pig cochlea. Eur Arch Otorhinolaryngol 256:323-329. Sunami K, Yamane H, Takayama M, Nakagawa T, Konishi K, Iguchi H (1999b): Cystine protects cochlear outer hair cells against glutamate toxicity. Acta Otolaryngol (Stockh) 119:671-673. Takumi Y, Matsubara A, Danbolt NC, Laake JH, Storm-Mathisen J, Usami S, Shinkawa H, Ottersen OP (1997): Discrete cellular and subcellular localization of glutamine synthetase and the glutamate transporter GLAST in the rat vestibular endorgan. Neuroscience 79:1137-1144. Takumi Y, Nagelhus EA, Eidet J, Matsubara A, Usami S, Shinkawa H, Nielsen S, Ottersen OP (1998): Select types of supporting cell in the inner ear express aquaporin-4 water channel protein. Eur J Neurosci 10:3584-3595. Takumi Y, Matsubara A, Laake JH, Ramirez-Leon V, Roberg B, Torgner I, Kvamme E, Usami S, Ottersen OP (1999a): Phosphate activated glutaminase is concentrated in mitochondria of sensory hair cells in rat inner ear: a high resolution immunogold study. J Neurocytol 28:223-237. Takumi Y, Ramirez-Leon V, Laake P, Rinvik E, Ottersen OP (1999b): Different modes of expression of AMPA and NMDA receptors in hippocampal synapses. Nat Neurosci 2:618-624. Tamarappoo BK, Raizada MK, Kilberg MS (1997): Identification of a system N-like Na(§ glutamine transport activity in rat brain neurons. J Neurochem 68:954-960. Usami S, Ottersen OP (1995): Differential cellular distribution of glutamate and glutamine in the rat vestibular endorgans: an immunocytochemical study. Brain Res 676:285-292. Usami S, Hozawa J, Tazawa M, Yoshihara T, Igarashi M, Thompson GC (1988): Immunocytochemical study of catecholaminergic innervation in the guinea pig cochlea. Acta Otolaryngol Suppl (Stockh) 447:36-45. Usami S, Osen KK, Zhang N, Ottersen OP (1992): Distribution of glutamate-like and glutamine-like immunoreactivities in the rat organ of Corti: a light microscopic and semiquantitative electron microscopic analysis with a note on the localization of aspartate. Exp Brain Res 91:1-11. Usami S, Matsubara A, Fujita S, Shinkawa H, Hayashi M (1995): NMDA (NMDAR1) and AMPA-type (GluR2/3) receptor subunits are expressed in the inner ear. Neuroreport 6:1161-1164. Yamashita H, Kawakami H, Zhang YX, Tanaka K, Nakamura S (1995): Neuroprotective mechanism of bromocriptine. Lancet 346:1305. Yamashita H, Kawakami H, Zhang YX, Tanaka K, Nakamura S (1998): Effect of amino acid ergot alkaloids on glutamate transport via human glutamate transporter hGluT-1. J Neurol Sci 155:31-36.

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CHAPTER X

A model glutamate synapse reticulospinal axon

the lamprey giant

O. SHUPLIAKOV AND L. BRODIN

1. INTRODUCTION Among the different types of fast-acting synapses, some exhibit unusual features which make it possible to study aspects of synaptic transmission that are normally inaccessible. Those synapses which have a large presynaptic element have been particularly valuable since they have permitted studies of the presynaptic machinery with direct methods. They include the squid giant synapse (Llin~is et al., 1992; Hunt et al., 1994), the goldfish giant bipolar terminal (Von Gersdorff and Matthews, 1999; see also Chapter 9 of this volume by Usami et al.), the 'synapse of Held' in the rat brainstem (Borst and Sakmann, 1996), and the lamprey giant reticulospinal synapse. In this chapter we will describe the latter synapse. The reticulospinal synapse utilizes glutamate as a neurotransmitter and has been used as a model system to determine the subcellular localization and uptake of glutamate in synaptic regions (Shupliakov et al., 1992, 1997b; Gundersen et al., 1995). It has also proved to be a powerful model in the analysis of molecular mechanisms in synaptic vesicle cycling (Pieribone et al., 1995; Shupliakov et al., 1997a; Gad et al., 1998; Ringstad et al., 1999).

2. THE LAMPREY RETICULOSPINAL S Y N A P S E -

AN OVERVIEW

The lamprey central nervous system has a similar organization as that in other vertebrates, although it lacks myelin and the number of neurons is comparatively low (Nieuwenhuys et al., 1998). The largest neurons in the lamprey CNS are the giant reticulospinal neurons or Mtiller cells. Their number is between 7 and 10 on either side of the brain depending on the criteria used to distinguish them from other reticulospinal neurons. The giant reticulospinal neurons (hereafter referred to as 'reticulospinal neurons') are divided into mesencephalic, isthmic, and bulbar neurons, based on the localization of the cell body (Nieuwenhuys et al., 1998). The reticulospinal neurons are part of the descending motor system with a primary function to transmit rapid motor commands, like postural signals and steering commands (Grillner et al., 1995). As their activity pattern is characterized by burst firing (Kasicki et al., 1989), the reticulospinal neurons can be classified as 'phasic' neurons (as opposed to 'tonic' neurons; Atwood and Wojtowicz, 1986; Brodin et al., 1997). Reticulospinal neurons form mixed electrotonic and chemical output synapses with motoneurons and different classes of interneurons along the spinal cord (Rovainen, 1974, 1979; Buchanan and Grillner, 1987; Grillner et al., 1995). The chemical synapses release glutamate, Handbook of Chemical Neuroanatomy, Vol. 18." Glutamate O.P. Ottersen and J. Storm-Mathisen, editors (~ 2000 Elsevier Science B.V. All rights reserved.

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acting at N-methyl-D-aspartate (NMDA) receptors, (RS)-alpha-amino-3-hydroxy-5-methyl4-isoxazolepropionic acid (AMPA) receptors (Buchanan et al., 1987), and metabotropic glutamate receptors (Krieger et al., 1996). Below we will describe the structure and function of release sites of this chemical synapse. Other aspects of the lamprey reticulospinal system have been reviewed elsewhere (Rovainen, 1979; Brodin et al., 1988; Grillner et al., 1995).

3. O R G A N I Z A T I O N OF T H E R E T I C U L O S P I N A L AXON The axons of the giant reticulospinal neurons project ipsilaterally and run in the ventromedial column throughout the length of the spinal cord. The only exception is the axon of the Mauthner cell, which crosses the midline at the brainstem level and runs in the contralateral dorsolateral column. The giant axons have a diameter ranging between 40 and 80 I~m, which makes them the fastest conducting axons in the animal. The conduction velocity is in the range of 3-7 m/s at temperatures of 8-10~ (Rovainen, 1979). Unlike most axons in the CNS of vertebrates, the reticulospinal axons do not ramify, but remain unbranched throughout their extent (Fig. 1A). The synaptic contact sites are established between the large axon stem and the postsynaptic neuron and are referred to as 'en passant' synapses (Fig. 1A,D). Hence, the release sites are distributed over the axonal plasmalemma directly exposed to the axoplasmic matrix. The release sites can be visualized in the living axon by microinjection of fluorescence-tagged antibodies to synaptic vesicle proteins (Pieribone et al., 1995). In such experiments (Fig. 1A-C), most release sites appear as small isolated spots without an evident pattern of organization within the axon. In some cases the spots are grouped together, and some of the spots appear larger, presumably reflecting multiple release sites. At the ultrastructural level the majority of the release sites (65-70%) appear as a single cluster of synaptic vesicles accumulated at a single active zone (Fig. 1D), which may contain specializations of pure chemical or mixed synapses (Pfenninger and Rovainen, 1974; Ringham, 1975; Christensen, 1976; Rovainen, 1979). The latter type thus contains gap junctions as well as active zones (Fig. 1D). Most of the synapses are established on dendritic shafts. A distinct class of interneuron, however, receives axo-somatic synapses (OS and LB unpublished). The majority of the postsynaptic cells receive multiple chemical or mixed synaptic contacts from a single axon. Therefore most of the EPSPs recorded in postsynaptic neurons are composed of two components, an electrotonic and a chemical component. The vesicle cluster and the active zone vary in size between individual synapses. Quantitative analysis has revealed correlations (1) between the total number of vesicles per synapse and the number of vesicles in the central section (i.e. through the midpoint of the active zone), and (2) between the number of synaptic vesicles in the central section and the length of the active zone in the central section (Fig. 1E,F; Shupliakov et al., 1995a). These correlations have made it possible to use the number of synaptic vesicles in the central section as an index of the number of vesicles in the whole cluster (Pieribone et al., 1995; Shupliakov et al., 1997a; Ringstad et al., 1999). It should be noted, however, that a smaller proportion of the synapses have a more complex organization which does not permit the use of the above correlations. For instance, analysis of serial ultrathin sections have demonstrated that closely located synapses may share one cluster of synaptic vesicles, and in addition unusually large active zones exceeding 2 txm in diameter are also present. The height of the synaptic vesicle cluster does not exceed 2 I~m, suggesting that this parameter is constrained. Single actiVe zones are surrounded by glial cell processes which surround the synaptic cleft like a collar and thereby provide a barrier to the extracellular space (Figs. 1D and 3). 274

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Fig. 1. Morphological organization of the reticulospinal synapse. (A) Schematic diagram showing the structural relation between a reticulospinal neuron and a spinal neuron in lamprey, and the experimental paradigm used to visualize active zones in the living giant axon (shown in B and C). An antibody against the synaptic vesicle-associated protein, synapsin, was labeled with Cy5 and pressure-injected into the axon. The distribution of fluorescently tagged antibodies was monitored with a CCD detector. Injection of the antibody into lamprey reticulospinal axons resulted in accumulation of fluorescence in spots, which are shown in B and C at different magnifications. The spots indicate the position of active zones (see also Pieribone et al., 1995). Scale bar in B, 25 Ixm for B and 15 Ixm for C. Inset in C is a cross-sectional confocal image of an injected axon which reveals that spots are localized to the inner surface of the axonal membrane (dashed line). Scale bar for the inset 2 t~m. (D) Electron micrograph of a reticulospinal synapse in a region of the axon outside the site of injection (indicated by a rectangle in A). Designations: sv, synaptic vesicle cluster; ax, axoplasmic matrix; gj, gap junction; g, glia; d, dendrite of a postsynaptic cell; arrowhead indicates the active zone. Scale bar, 0.2 t~m. Note the presence of specializations of both an electrical and a chemical synapse in the same intracellular contact. (E) Correlation between the total number of synaptic vesicles per synapse and the number of vesicles in the center section of reticulospinal axon synapses (correlation coefficient; r = 0.91). (F) Correlation between the number of synaptic vesicles in the center section and the length of the active zone (r -- 0.90). B and C, modified from Pieribone et al. (1995): Nature 375:493-497, with permission; copyright Macmillan Magazines Ltd. E and F, reprinted from Shupliakov et al. (1995): Eur J Neurosci 7:1111-1116, with permission.

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4. SYNAPTIC LOCALIZATION OF GLUTAMATE AND RELATED AMINO ACIDS The organization of the reticulospinal axons has permitted detailed analysis of the distribution of glutamate and other amino acids in different subcompartments of the synapse. Studies with quantitative post-embedding methods (Ottersen, 1989) have shown that the level of glutamate labeling is about 20 times higher over the synaptic vesicle cluster as compared to the surrounding axoplasmic matrix (Fig. 2A-C; Shupliakov et al., 1992). By comparing the particle density with that in co-processed test conjugates, the glutamate concentration in the synaptic vesicle cluster was estimated to be above 30 mM (Fig. 2B), indicating an intravesicular concentration of more than 60 mM (Shupliakov et al., 1992). As some glutamate may be lost during fixation (Ottersen, 1989) the glutamate concentration is likely to be even higher in vivo. Other amino acids, such as aspartate, homocysteate, and glycine were not found to be accumulated in this region (Shupliakov et al., 1992, 1996). To localize the site of glutamate uptake in synaptic regions of the giant synapse, the lamprey spinal cord preparation was incubated with the metabolically inert transporter substrate D-aspartate, followed by immunogold labeling with D-aspartate antibodies (Gundersen et al., 1995). The most intense D-aspartate labeling was detected in astroglial processes (Fig. 3A), which surround the synapses. The uptake into neuronal elements (both pre- and postsynaptic) was found to be limited. The results of these experiments also indicated that D-aspartate is not taken up into synaptic vesicles in the intact synapse. Thus, prolonged incubation with a high concentration of O-aspartate (500 IxM; 10 h; i.e. near the Km value established for vesicular glutamate transporters; Tabb and Ueda, 1991) did not result in any significant labeling over synaptic vesicle clusters, even when combined with electrical stimulation (Gundersen et al., 1995). Although the uptake of an exogenously applied glutamate analog is not directly equivalent to the uptake of synaptically released glutamate, these findings indicate that the glial processes surrounding the synapses play an important role in clearing glutamate from the synaptic region. The effective uptake of glutamate into glial processes is consistent with the glutamateglutamine cycle hypothesis, which implies that glutamate is shuttled between nerve terminals and glia (see Chapter 1 by Broman et al. and Chapter 7 by Kaneko in this volume). The released glutamate is thought to be converted to glutamine in the latter, and reconverted to glutamate in the former. The levels of glutamine are high in glial processes (Fig. 2D) and low in presynaptic mitochondria and axoplasm, which also agrees with this hypothesis (Shupliakov et al., 1997b). When the levels of amino acids in the phasic reticulospinal axon were compared with those in a tonic glutamatergic axon (the sensory dorsal column axons), some notable differences were observed. First, the level of glutamate labeling over axoplasmic matrix and presynaptic mitochondria was found to be about 4 times higher in the tonic axons as compared to that in the reticulospinal axons (Fig. 2C; Shupliakov et al., 1997b). Second, after incubation with exogenous D-aspartate, the labeling was significantly higher in glial processes around dorsal column synapses as compared to those around reticulospinal synapses (Gundersen et al., 1995). Third, the level of glutamine in the glial processes showed a corresponding difference (Fig. 2D; Shupliakov et al., 1997b). These observations indicate that the pool of transmitter glutamate is larger and more effectively circulated at a tonic synapse as compared to the phasic reticulospinal synapse, which appears physiologically relevant. This conclusion has been supported by studies of phasic and tonic glutamatergic synapses in the crayfish neuromuscular system (Shupliakov et al., 1995b).

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Fig. 2. Glutamate and glutamine immunoreactivity in the reticulospinal synapse. (A) Electron micrograph of a reticulospinal synapse stained with an antiserum to fixed glutamate using the post-embedding immunogold technique. All designations as in Fig. 1. The area containing synaptic vesicles (sv) displays a higher density of gold particles compared to axoplasmic matrix (ax) or mitochrondria (m), dendrites (d) and glial elements (g). (In ax, cross-sectioned filaments and microtubules should not be confused with gold particles.) Insets in A show 'test protein-glutaraldehyde-amino acid conjugates' with glutamate (GLU), aspartate (ASP), and glutamine (GLN), respectively, incubated along with tissue sections to monitor the specificity of the staining reaction. 'None' represents conjugates made by reacting a brain macromolecule extract with glutaraldehyde without addition of amino acids (see e.g. Ottersen, 1989). Note the specific accumulation of 15 nm gold particles over the glutamate conjugate. Scale bar, 0.2 I~m. (B) Relationship between the concentration of fixed glutamate and the density of gold particles in test conjugates used to estimate the concentration of glutamate in co-processed tissue sections. The diagram shows the relation between the gold particle density and the concentration of fixed glutamate in the test conjugates. The relationship was linear within the examined concentration range. The bars represent SEM. The circle on the line indicates the concentration of fixed glutamate which corresponds to the density of gold particles present over the synaptic vesicle cluster in the synapse shown above. (C, D) Histograms showing the distribution of glutamate (C) and glutamine (D) labeling over different cell compartments of elements composing reticulospinal and dorsal column synapses, respectively. Bars represent average densities (• of gold particles over the compartments. The densities (particles/l~m 2) represent arbitrary units, i.e. they have not been corrected for the different labeling efficiencies for glutamine and glutamate. Background over tissue-free resin (<2 particles/l~m 2) was subtracted. Asterisks indicate statistically significant differences (see Shupliakov et al., 1997a,b). A and B, reprinted from Brodin et al. (1994): Adv Second Messenger Phosphoprotein Res 29:205-221; C and D, reprinted from Shupliakov et al. (1997b)" Neuroscience 77:1201-1212, with permission.

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Fig. 3. Organization of pre- and postsynaptic elements and glial cell processes in reticulospinal synapses. (A, B) Three-dimensional reconstructions of a dendritic shaft (d) receiving synapses from a reticulospinal axon (ax). The sites of synaptic contact are indicated with arrows. The surface of the axon was drawn from one section in the middle of the series of sections (synaptic vesicle clusters were not included). The glial processes (g) occupy the entire space (marked with asterisks) in between the dendritic shafts and the surface of the axons. A, shows the reconstruction as seen vertical to the plane of cutting (i.e. a view from rostral to caudal). B, shows 90 ~ rotation giving a view from dorsal to ventral. The glial profiles occupy the entire shaded area. Note that the shaded area of glial processes surrounds the areas of synaptic contacts between the axon and the dendrite (dark areas indicated with arrows). The position of the dendritic shaft, which is obscured by glial cell processes, is indicated with dashed lines. The surface of the axon was not included in this projection. Scale bar, 1 lxm. (C) Electron micrograph of a reticulospinal synapse from a specimen incubated in Ringer solution containing 50 IxM D-aspartate for 1 h. Gold particles indicate an accumulation of D-aspartate immunoreactivity over glial cell processes surrounding the synapse. Scale bar, 0.2 ~tm. Other designations as in Fig. 1. (D, E) Quantitative determination of D-Asp immunolabeling in various tissue compartments in specimens incubated with 50 IxM or 500 IxM D-Asp for 1 h. The values are mean numbers of gold particles/Ixm 2 -t- SEM corrected for background immunoreactivity over empty resin (0.5 particles/Ixm2). Asterisks indicate that the density of D-Asp immunogold particles in glial processes around synapses at 50 IxM (n = 65) and 500 IxM (n = 20) D-Asp is significantly different from the density in other tissue compartments (p < 0.001). Values in other compartments were not significantly different from the background labeling. Reprinted from Gundersen et al. (1995): J Neurosci 15:4417-4428, with permission.

Lamprey giant reticulospinal axon

Ch. X

5. SYNAPTIC VESICLE POOLS The synaptic vesicle cluster at a reticulospinal synapse appears as a dense homogeneous mass of synaptic vesicles. Studies in which the synaptic vesicle-associated proteins, synapsins, were perturbed suggest, however, that the vesicle cluster consists of two functionally distinct pools (Pieribone et al., 1995). When synapsin antibodies (directed to domain E of synapsins Ia, IIa) were microinjected into the living axon, the distal part of the vesicle cluster was disrupted (Fig. 4A,B; Pieribone et al., 1995). In contrast, the small pool located adjacent to the plasma membrane remained intact. By performing immunogold labeling, synapsins were found to be primarily concentrated in the major distal part of the vesicle cluster (Fig. 4C). Thus, the synaptic vesicle cluster can be divided into a large distal pool which depends on synapsins, and a small proximal synapsin-independent pool. The factors involved in the organization of the synapsin-independent pool are as yet unknown. It is interesting to note, however, that labeling with phosphotungstic acid (Fig. 4D) reveals a matrix of filaments extending from the plasma membrane which has a distribution overlapping with the synapsin-independent vesicle pool (Brodin et al., 1997). Recent findings that a novel set of proteins, including Bassoon, Piccolo/Aczonin, and Rim are accumulated in this area in mammalian synapses suggest that specific molecular mechanisms are involved in the regulation of the synaptic vesicle pool adjacent to the active zone (Cases-Langhoff et al., 1996; Wang et al., 1997; Tom-Dieck et al., 1998; Wang et al., 1999). Data from electrophysiological studies suggest that the two vesicle pools play distinct functional roles. Following microinjection of synapsin antibodies, synaptic transmission at low frequencies remained unaffected, indicating that the synapsin-independent pool is sufficient to support basal release (Pieribone et al., 1995). During high-frequency stimulation a marked depression of the synaptic response was uncovered (Fig. 5), which was paralleled by an almost complete depletion of synaptic vesicles at release sites. The synapsin-dependent vesicle pool thus appears to be required to sustain release during periods of high-frequency firing. Studies of genetically modified mice indicate that synapsins play a similar role in mammals. The phenotype of synapsin-deficient mice thus includes disrupted synaptic vesicle clusters and an enhanced synaptic depression during high-frequency stimulation (Li et al., 1995; Rosahl et al., 1995; Takei et al., 1995).

6. P R E S Y N A P T I C Ca 2+ C H A N N E L S

The structure of the reticulospinal axon, with synaptic release sites scattered at low density over a large area of plasma membrane, implies that the presynaptic Ca 2+ current only represents a tiny fraction of the total ionic currents over the axonal membrane. Consequently, the presynaptic Ca 2+ current is difficult to record in this system. However, after blockade of Na + and K + channels, and after addition of high concentrations of BaCI2, Ba 2+ potentials can be recorded with a two-electrode current clamp (MacVicar and Llimis, 1985; Shupliakov et al., 1995a). These Ba 2+ potentials provide a means to monitor the function of presynaptic Ca 2+ channels during experimental manipulation of the release machinery (Shupliakov et al., 1995a; L6w et al., 1999). Although it has not yet been directly confirmed that these Ba 2+ potentials reflect the Ca 2+ current flowing at active zones, the close association of Ca 2+ channels with release sites in other systems favor this possibility (Llimis et al., 1992; Borst and Sakmann, 1996; Von Gersdorff and Matthews, 1999). The pharmacology of the presynaptic Ca 2+ channels in the reticulospinal synapse has been 279

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F/g. 4. Disruption of the synapsin-dependent vesicle pool by presynaptic microinjection of synapsin antibodies in the lamprey reticulospinal synapse. (A) Electron micrograph of a 'control' synapse. (B) A synapse in an axon injected with synapsin antibodies. The axon was lightly stimulated (1 Hz for 12 min) and allowed to rest for 90 min before fixation. Note that a narrow rim of vesicles remains in the antibody-injected synapse. (C) Immunogold staining of a reticulospinal synapse with synapsin antibodies. Note that the vesicles adjacent to the presynaptic membrane are almost devoid of gold particles. (D) Visualization of the filamentous cytomatrix that overlaps with the synaptic vesicle pool that remains after perturbation of synapsins. The electron micrograph shows a synapse in a normal axon (i.e. no microinjection or stimulation had been performed) stained with phosphotungstic acid (Gustafsson et al., 1996). The filamentous cytomatrix (arrowheads) at the presynaptic membrane is visible, but not the synaptic vesicle cluster. Designations as in Fig. 1. Scale bar, 0.2 Ltm. Reprinted from Brodin et al. (1995): Eur J Neurosci 9:2503-2511, with permission.

studied by monitoring the effect of channel inhibitors on the reticulospinal EPSP (Krieger et al., 1999). The N-type Ca 2+ channel blocker, omega-conotoxin GVIA, was found to induce a large decrease of the reticulospinal EPSP, whereas the P/Q-type calcium channel blocker, omega-agatoxin IVA, produced a smaller inhibition. Combined application of the two toxins produced a strong depression of the EPSP. It did not cause a complete blockade, however, indicating that calcium channels insensitive to these toxins (R-type) are also involved. Thus presynaptic calcium influx in reticulospinal axons appears to be mediated through N-, P/Q280

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Fig. 5. Impairment of high-frequency synaptic transmission after microinjection of synapsin antibodies. (A) Control responses recorded before microinjection of synapsin antibodies at 18 Hz. The plot shows amplitudes of EPSPs evoked in a spinal neuron from a lamprey reticulospinal axon. The traces show averages of EPSPs during the two recording periods (1 and 2, respectively) (B) Responses recorded after microinjection of synapsin antibodies. Note the enhanced depression after high-frequency stimulation. (C, D) Changes in synaptic ultrastructure induced by high-frequency stimulation. (C) A synapse in an 'uninjected' control axon. (D) A synapse within an axon injected with synapsin antibodies. Both axons were from the same spinal cord, which was stimulated at 18 Hz for 6 rain immediately prior to fixation. Note the depletion of synaptic vesicles at release sites. Scale bar, 0.2 I~m. Modified from Pieribone et al. (1995): N a t u r e 3 7 5 : 4 9 3 - 4 9 7 , with permission; copyright 1995 Macmillan Magazines Ltd.

and R-type channels, with N-type channels playing the major role. L-type channels are unlikely to be involved, as agonists and antagonists of these channels have been found to be ineffective (Krieger et al., 1999).

7. PRESYNAPTIC MODULATION OF TRANSMITTER RELEASE The reticulospinal axons are surrounded by a dense plexus of 5-HT-containing fibers (Grillner et al., 1995). Application of 5-HT produces a strong inhibition of the reticulospinal EPSE Two lines of evidence indicate that this effect is due to a presynaptic modulation of release. First, 5-HT does not reduce the response to exogenous glutamate in the postsynaptic neuron (Buchanan and Grillner, 1991). Second, the depletion of synaptic vesicle clusters induced by 281

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......iX-mv Fig. 6. Activation of two distinct types of metabotropic glutamate receptor depresses monosynaptic reticulospinal EPSPs. Effect of (S)-2-amino-2-methyl-4-phosphonobutanoic acid (MAP4) on (1S,3R)- 1-aminocyclopentane-l,3-dicarboxylic acid/(1S,3R)-ACPD/- and L(+)-2-amino-4-phosphonobutyric acid/L-AP4/-induced depression of the EPSP. (A) Effect of (1S,3R)-ACPD (10 IxM) and L-AP4 (50 [tM) on the amplitude of the reticulospinal-evoked EPSP was tested before and during application of MAP4 (1 mM). Monosynaptic EPSPs during periods marked a-e are shown in B and C. (B) Depression of the monosynaptic EPSPs by (1S,3R)-ACPD was not antagonized by MAP4. Inset: input resistance of the recorded neuron did not change during application of (1S,3R)-ACPD. (C) L-AP4-induced decrease of the monosynaptic EPSP was reduced by MAP4. Inset: L-AP4 did not affect the input resistance of the postsynaptic neuron. Reprinted from Krieger et al. (1996): J Neurophysiol 76:3834-3841, with permission.

intense action potential stimulation (see below; Wickelgren et al., 1985) is prevented by 5-HT application, which implies that the rate of exocytosis is reduced (Shupliakov et al., 1995a). The 5-HT-mediated presynaptic inhibition does not appear to be due to a reduced presynaptic Ca 2+ influx, as axonal Ba 2+ spikes are unaffected by 5-HT (Shupliakov et al., 1995a). The reticulospinal EPSP can also be modulated via metabotropic glutamate receptors (Fig. 6). The mGluR agonists (1S,3R)- 1-aminocyclopentane- 1,3-dicarboxylic acid [(1S,3R)ACPD] and L(+)-2-amino-4-phosphonobutyric acid (L-AP4) were both found to reduce the amplitude of the reticulospinal EPSP, without affecting the amplitude of postsynaptic AMPA-induced depolarizations (Krieger et al., 1996). The mGluR antagonist alpha-methylL-AP4 blocked the depression induced by L-AP4 but not that induced by (1S,3R)-ACPD. Furthermore, the effects of co-application of (1S,3R)-ACPD and L-AP4 were additive. Thus at least two types of pharmacologically distinct presynaptic mGluRs appear to be present (Krieger et al., 1996). It is possible that these presynaptic mGluRs serve as glutamatergic autoreceptors, which provide a negative feedback to the release machinery. The above results were obtained in experiments with sharp electrodes in adult lampreys. In a study with whole282

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Fig. 7. Reversible trapping of synaptic vesicle membrane in the plasma membrane in reticulospinal synapses. (A) Electron micrograph of a lamprey reticulospinal synapse stimulated with action potentials at 20 Hz for 20 min and then incubated for 90 min in Cae+-free solution with 10 mM EGTA. Note the reduction in the number of synaptic vesicles and the presence of large membrane expansions compared to an unstimulated synapse (inset). (B) Activation of clathrin-mediated endocytosis in reticulospinal synapses by addition of Cae+-containing extracellular solution. Spinal cord preparations were stimulated at 20 Hz for 20 min, incubated for 90 min in Cae+-free solution, and then incubated in Cae+-containing solution (2.6 mM) for 120 s. Electron micrograph of a synapse shows the appearance of coated pits (arrows) lateral to the active zone. Designations as in Fig. 1. Scale bar, 0.2 Ixm. Modified from Gad et al. (1998): Neuron 21:607-616, with permission; copyright is held by Cell Press.

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cell recording in larval lampreys, Cochilla and Alford (1998) found that application of either ACPD or L-AP4 causes a reduction of the presynaptic spike amplitude due to activation of a K + conductance. Another mGluR agonist (RS)-3,5-dihydroxyphenylalanine (DHPG), was found to enhance the frequency-dependent facilitation of reticulospinal EPSPs. This effect was blocked by ryanodine suggesting that DHPG acted by enhancing Ca 2+ release from intracellular stores (Cochilla and Alford, 1998).

8. SYNAPTIC VESICLE RECYCLING When a reticulospinal axon is subjected to action-potential stimulation at a moderate rate (up to 5 Hz for 30 min), synaptic transmission can be sustained for long periods with little reduction in the size of synaptic vesicle clusters (Shupliakov et al., 1997a). Thus, at this rate of activity, the synaptic vesicle pools can be maintained by recycling, which involves clathrin-mediated endocytosis (see below). During brief periods of high-frequency stimulation, vesicles can be drawn from the synapsin-dependent vesicle pool (see above; Pieribone et al., 1995), thus permitting the rate of exocytosis to intermittently exceed the rate of recycling. If high-frequency stimulation is continued for longer periods, however, a progressive depletion of vesicles occurs, and extensive plasma membrane invaginations appear around the active zones as a result of the incorporation of synaptic vesicle membrane (Wickelgren et al., 1985). Such massive vesicle depletion is most likely an unphysiological condition (i.e. the phasic reticulospinal synapse is not normally exposed to tonic activity), but it can be used experimentally to study synaptic vesicle endocytosis. Gad et al. (1998) used a high-frequency stimulation protocol to test if synaptic vesicle endocytosis can be dissociated from CaZ+-evoked exocytosis. Following depleting stimulation, the rapid removal of extracellular Ca 2+ was found to arrest the recovery of the vesicle pool, and the plasma membrane remained expanded (Fig. 7A). By applying a low concentration of Ca 2+ (11 txM), the recovery process could be activated (without any action-potential stimulation) after more than 1 h. Thus, synaptic vesicle endocytosis can be temporally separated from CaZ+-evoked exocytosis. These experiments also showed that the endocytosis of synaptic vesicles requires a low level of Ca 2+. The activation of vesicle recycling in the above experiments was associated with a massive accumulation of clathrin-coated pits in the plasma membrane around active zones (Fig. 7B). At early times (10-20 s) after addition of Ca 2+, early stages of coated pits (i.e. shallow coated pits) were relatively more abundant, whereas at later times (2 min), late stages (i.e. invaginated coated pits with narrow necks) predominated. Synaptic vesicle recycling under conditions of low-frequency stimulation also appears to be predominantly or exclusively mediated by clathrin-mediated endocytosis. In axons maintained at rest, clathrin-coated pits

Fig. 8. Effects of disruption of endophilin and amphiphysin interactions on clathrin-mediated endocytosis at the reticulospinal synapse. (A) Electron micrograph of the lateral side of the active zone in a control synapse stimulated at 5 Hz. Note the presence of clathrin-coated pits with different shapes. (B) Electron micrograph of the comparable area of a synapse in an axon that was stimulated at 5 Hz for 30 min after injection of endophilin antibodies. Note the 'pocket-like' membrane expansions (arrows) at the margin of the synaptic area and the appearance of numerous 'shallow' coated pits (arrows). (C) A synapse in an axon which was stimulated at 0.2 Hz for 30 min after injection of a fusion protein containing the SH3 domain of amphiphysin linked to GST. Note the accumulation of constricted coated pits around the active zone. Scale bar, 0.2 gm. B, modified from Ringstad et al. (1999), Neuron 24, 143-154, with permission; copyright is held by Cell Press. C, modified from Shupliakov et al. (1997a): Science 276:259-263, with permission; copyright 1997 AAAS.

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are virtually absent at synaptic release sites. Following action-potential stimulation at low to moderate rates, a small but consistent number of coated pits appear in the plasma membrane around active zones (Shupliakov et al., 1997a). When proteins in the endocytic machinery are perturbed, however, a massive accumulation of clathrin-coated pits occurs in response to stimulation (see below; Shupliakov et al., 1997a; Ringstad et al., 1999).

9. MOLECULAR MECHANISMS IN SYNAPTIC VESICLE ENDOCYTOSIS The giant reticulospinal synapse has several features which makes it a suitable model to study the molecular mechanisms underlying clathrin-mediated synaptic vesicle endocytosis. First of all, the size of the reticulospinal axon permits presynaptic microinjections (see Fig. 1). Moreover, the organization of the release sites, with single active zones surrounded by an almost flat axonal membrane greatly facilitates electron microscopic analysis of endocytic intermediates around the synapses. Another advantage is that the rate of spontaneous release is low, which makes it possible to verify that the effect of a microinjected compound depends on vesicle cycling (by comparing the effect of an injected compound on the resting synapse with that in the stimulated synapse). By combining microinjections with electron microscopic analysis, it has been possible to link different proteins implicated in endocytosis (for reviews see: Cremona and De Camilli, 1997; Schmidt et al., 1999) to distinct endocytic intermediates in the intact synapse. For example, the endocytic protein, endophilin (Cremona and De Camilli, 1997; Schmidt et al., 1999), has been linked to an early step in the formation of clathrin-coated pits. Microinjection of anti-endophilin antibodies, followed by stimulation, led to a massive accumulation of shallow clathrin-coated pits studies around the release sites (Fig. 8; Ringstad et al., 1999). This indicates that endophilin plays an essential role in the invagination of the clathrin-coated pit. It is also possible to study the role of protein-protein interactions in endocytosis. Shupliakov et al. (1997a) used the reticulospinal synapse to examine the role of the two endocytic proteins, dynamin and amphiphysin (Cremona and De Camilli, 1997). Microinjection of proteins or peptides which inhibit the binding of the SH3 domain of amphiphysin to a specific binding site in the proline-rich domain of dynamin caused a stimulus-dependent accumulation of deeply invaginated clathrin-coated pits with a narrow constricted neck (Fig. 8C). These data suggest that the amphiphysin-dynamin interaction is essential in the process mediating fission of the neck of the coated pit. The above examples illustrate that detailed information about discrete molecular processes in endocytosis can be obtained by combining microinjections with electron microscopic analysis at the giant reticulospinal synapse.

10. CONCLUSIONS Central nerve terminals are heterogeneous with regard to structure, function and molecular composition. This diversity adds complexity to the function of neural circuits. From the experimental point-of-view, the heterogeneity of nerve terminals provide neuroscientists with a variety of models suited to address different aspects of synaptic function. The giant reticulospinal neuron in the lamprey has a very large unbranched axon with isolated active zones. These unique features make the glutamatergic reticulospinal axon one of the most powerful in vivo models available to study molecular mechanisms in synaptic vesicle cycling. 286

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11. REFERENCES Atwood HL, Wojtowicz JM (1986): Short-term and long-term plasticity and physiological differentiation of crustacean motor synapses. Int Rev Neurobiol 28:275-362. Borst JGG, Sakmann B (1996): Calcium influx and transmitter release in a fast CNS synapse. Nature 383:431-434. Brodin L, Grillner S, Dubuc R, Ohta Y, Kasicki S, H6kfelt T (1988): Reticulospinal neurons in lamprey: transmitters, synaptic interactions and their role during locomotion. Arch Ital Biol 126:317-345. Brodin L, L6w P, Gad H, Gustafsson J, Pieribone V, Shupliakov O (1997): Sustained neurotransmitter release: new molecular clues. Eur J Neurosci 9:2503-2511. Buchanan JT, Grillner S (1987): Newly identified 'glutamate interneurons' and their role in locomotion in the lamprey spinal cord. Science 236:312-314. Buchanan JT, Grillner S (1991): 5-Hydroxytryptamine depresses reticulospinal excitatory postsynaptic potentials in motoneurons of the lamprey. Neurosci Lett 122:71-74. Buchanan JT, Brodin L, Dale N, Grillner S (1987): Reticulospinal neurones activate excitatory amino acid receptors. Brain Res 408:321-325. Cases-Langhoff C, Voss B, Garner AM, Appeltauer U, Takei K, Kindler S, Veh RW, De Camilli P, Gundelfinger ED, Garner CC (1996): Piccolo, a novel 420 kDa protein associated with the presynaptic cytomatrix. Eur J Cell Biol 69:214-223. Christensen BN (1976): Morphological correlates of synaptic transmission in lamprey spinal cord. J Neurophysiol 39:197-212. Cochilla AJ, Alford S (1998): Metabotropic glutamate receptor-mediated control of neurotransmitter release. Neuron 20:1007-1016. Cremona O, De Camilli P (1997): Synaptic vesicle endocytosis. Curr Opin Neurobiol 7:323-330. Gad H, L6w P, Zotova E, Brodin L, Shupliakov O (1998): Temporal dissociation between Ca2+-induced synaptic vesicle exocytosis and clathrin-mediated endocytosis in a central synapse. Neuron 21:667-677. Grillner S, Deliagina T, Ekeberg O, E1 Manira A, Hill RH, Lansner A, Orlovsky GN, Wall6n P (1995): Neural networks that co-ordinate locomotion and body orientation in lamprey. Trends Neurosci 18:270-279. Gundersen V, Shupliakov O, Brodin L, Ottersen OP, Storm-Mathisen J (1995): Quantification of excitatory amino acid uptake at intact glutamatergic synapses by immunohistochemistry of exogenous D-aspartate. J Neurosci 15:4417-4428. Gustafsson J, Shupliakov O, Brodin L (1996): Electronmicroscopic visualization of structures controlling the "promimal pool" of synaptic veiscles in the lamprey reticulospinal synapse. Abstr Scandanivian Physiological Meeting, Stockholm, p. 50. Heuser JE (1989): The role of coated vesicles in recycling of synaptic vesicle membrane. Cell Biol Int Rep 13:1063-1076. Hunt JM, Bommert K, Charlton MP, Kistner A, Habermann E, Augustine GJ, Betz H (1994): A post-docking role for synaptobrevin in synaptic vesicle fusion. Neuron 12:1269-1279. Kasicki S, Grillner S, Ohta Y, Dubuc R, Brodin L (1989): Phasic modulation of reticulospinal neurones during fictive locomotion and other types of spinal motor activity in lamprey. Brain Res 484:203-216. Krieger P, El Manira A, Grillner S (1996): Activation of pharmacologically distinct metabotropic glutamate receptors depresses reticulospinal-evoked monosynaptic EPSPs in the lamprey spinal cord. J Neurophysiol 76:3834-3841. Krieger P, Buschges A, E1 Manira A (1999): Calcium channels involved in synaptic transmission from reticulospinal axons in lamprey. J Neurophysiol 81:1699-1705. Li L, Chin L-S, Shupliakov O, Brodin L, Sihra TS, Hvalby OS, Jensen V, Zheng D, McNamara J, Greengard P, Andersen P (1995): Impairment of synaptic vesicle clustering and of synaptic transmission, and increased seizure propensity in synapsin I-deficient mice. Proc Natl Acad Sci USA 92:9235-9239. Llimis R, Sugimori M, Silver RB (1992): Microdomains of high calcium concentration in a presynaptic terminal. Science 256:677-679. L6w P, Norlin T, Risinger C, Larhammar D, Pieribone VA, Shupliakov O, Brodin L (1999): Effects of SNAP-25 antibodies on neurotransmitter release and synaptic vesicle organization in a vertebrate giant synapse. Eur J Cell Biol 78:787-793. MacVicar BA, Llin~is R (1985): Barium action potentials in regenerating axons of the lamprey spinal cord. J Neurosci Res 13:323-335. Nieuwenhuys R, Ten Donkelaar HJ, Nicholson C (1998): The Central Nervous System of Vertebrates, Vol. 1. Berlin: Springer. Ottersen OP (1989): Quantitative electron microscopic localization of neuroactive amino acids. Anat Embryol 180:1-15.

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Pfenninger KH, Rovainen CM (1974): Stimulation- and calcium-dependence of vesicle attachment sites in the presynaptic membrane: a freeze-cleave study on the lamprey spinal cord. Brain Res 72:1-23. Pieribone VA, Shupliakov O, Brodin L, Hilfiker-Rothenfluh S, Czernik AJ, Greengard P (1995): Distinct pools of synaptic vesicles in neurotransmitter release. Nature 375:493-497. Ringham GL (1975): Localization and electrical characteristics of a giant synapse in the spinal cord of the lamprey. J Physiol 251:395-407. Ringstad N, Gad H, L6w R Takei K, Brodin L, Shupliakov O, De Camilli P (1999): Endophilin/SH3p4 is required for the transition from early to late stages in clathrin-mediated synaptic vesicle endocytosis. Neuron 24:143-154. Rosahl TW, Spillane D, Missler M, Herz J, Selig DK, Wolff JR, Hammer RE, Malenka RC, Stidhof TC (1995): Essential functions of synapsins ! and II in synaptic vesicle regulation. Nature 375:488-493. Rovainen CM (1974): Synaptic interactions of reticulospinal neurons and nerve cells in the spinal cord of the sea lamprey. J Comp Neurol 154:207-223. Rovainen CM (1979): Neurobiology of lampreys. Physiol Rev 59:1007-1077. Schmidt A, Wolde M, Thiele C, Fest W, Kratzin H, Podtelejnikov AV, Witke W, Huttner WB, S61ing HD (1999): Endophilin I mediates synaptic vesicle formation by transfer of arachidonate to lysophosphatidic acid. Nature 401:133-141. Shupliakov O, Brodin L, Cullheim S, Ottersen OR Storm-Mathisen J (1992): Immunogold quantification of glutamate in two types of excitatory synapse with different firing patterns. J Neurosci 12:3789-3803. Shupliakov O, Pieribone V, Gad H, Brodin L (1995a): Synaptic vesicle depletion in reticulospinal axons is reduced by 5-HT: direct evidence for presynaptic inhibition of glutamatergic transmission. Eur J Neurosci 7:1111-1116. Shupliakov O, Atwood HL, Storm-Mathisen J, Ottersen OR Brodin L (1995b): Presynaptic glutamate levels in tonic and phasic excitatory motor axons correlate with properties of synaptic release. J Neurosci 15:7168-7180. Shupliakov O, Pieribone VA, Gad H, Brodin L (1996): Presynaptic mechanisms in central synaptic transmission: 'biochemistry' of an intact glutamatergic synapse. Acta Physiol Scand 157:369-379. Shupliakov O, L6w R Grabs D, Gad H, Chen H, David C, Takei K, De Camilli R Brodin L (1997a): Synaptic vesicle endocytosis impaired by disruption of dynamin-SH3 domain interactions. Science 276:259-263. Shupliakov O, Storm-Mathisen J, Ottersen OR Brodin L (1997b): Glial and neuronal glutamine pools at glutamatergic synapses with distinct properties. Neuroscience 77:201-1212. Tabb JS, Ueda T (1991): Phylogenetic studies on the synaptic vesicle glutamate transport system. J Neurosci 11:1822-1828. Takei Y, Harada A, Takeda K, Kobayashi K, Terada K, Noda T, Takahashi T, Hirokawa N (1995): Synapsin I deficiency results in the structural change in the presynaptic terminals in the murine nervous system. J Cell Biol 131:1789-1800. Tom-Dieck S, Sanmarti-Vila L, Langnaese K, Richter K, Kindler S, Soyke A, Wex H, Smalla KH, Kampf U, Franzer JT, Stumm M, Garner CC, Gundelfinger ED (1998): Bassoon, a novel zinc-finger CAG/glutamine-repeat protein selectively localized at the active zone of presynaptic nerve terminals. J Cell Biol 142:499-509. Von Gersdorff H, Matthews G (1999): Electrophysiology of synaptic vesicle cycling. Annu Rev Physiol 61:725752. Wang XL, Kibschull M, Laue MM, Lichte B, Petrasch-Parwez E, Kilimann MW (1999): Aczonin, a 550-kD putative scaffolding protein of presynaptic active zones, shares homology regions with rim and bassoon and binds profilin. J Cell Biol 147:151-162. Wang Y, Okamoto M, Schmitz F, Hofmann K, Sudhof TC (1997): Rim is a putative Rab3 effector in regulating synaptic-vesicle fusion. Nature 388:593-598. Wickelgren WO, Leonard JR Grimes MJ, Clark RD (1985): Ultrastructural correlates of transmitter release in presynaptic areas of lamprey reticulospinal axons. J Neurosci 5:1188-1201.

288

Subject Index

t~-actinin 189 adrenal gland 156 AKAP 190 amygdala 26, 152

release 47 calcium dependence 49 synaptic vesicles 51 vasodilatation 54

AMPA receptor binding protein (ABP) 191

D-aspartate

AMPA receptors 99 adrenal gland 156 anchoring 171 calcium permeability 101, 136 cerebellum 132, 153 cochlear nuclei 155, 159 cytoplasmic distribution 168 flip/flop isoforms 122 hippocampus 121, 149 hypothalamus 152 inner ear 262 interaction with NSF 192 mRNA distribution 101 neocortex 115, 149 olfactory bulb 149 Q/R site editing 136 retina 112, 157 RNA splicing 122 spinal cord 135, 155 substantia nigra 153 synaptic distribution 162, 265 targeting 171, 190 thalamus 152 transport 168 turnover 193 amphiphysin 286 arachidonic acid 242 aspartate 45 colocalization 52 heteroexchange 50 immunocytochemistry 51 in hypoglycemia 46 LTP 53 nerve terminal localization 46

aspartate aminotransferase

retrograde tracer 2,29, 276 5, 46, 221,223 localization 223

basal ganglia 25, 55, 126, 151 Bassoon 279 calcium channels presynaptic 279 pharmacology 280

calmodulin 189 calmodulin dependent protein kinase II (CaMKII) 187

calyces of Held 18 cerebellum 20, 55, 129 climbing fibers 20, 55, 166 mossy fibers 20, 161, 215 parallel fibers 22, 161 cerebral cortex 28, 113

Chinese restaurant syndrome 157

citron 187 clathrin 284 cochlea 56 cochlear nuclei 18 eortieothalamic projection 23 CRIPT 164

delta glutamate receptors 101,161 anchoring 193

dihydrokainic acid 49 Dig 185

dynamin 286 Eph receptor tyrosine kinases 192

Fyn tyrosine kinase 188 GABA synthesis 223

GKAP 188 glucose glutamate precursor 3

glucose transport GLUT1 3 GLUT3 3

glutamate degradation 203 immunocytochemistry 2,11 release 262, 279 modulation 281 synaptic localization 276 synthesis 3,203,219 time course in synaptic cleft 243 turnover 10 uptake 10 chloride conductance 233,243 mechanisms 232 in nerve terminals 232 regulation 241 role in synaptic transmission 243 stoichiometry 233 vesicular uptake 8 glutamate dehydrogenase 220 glutamate transporters 231 developmental changes 240 concentration 240 localization 241 EAAT 1 (GLAST) 219 concentration 237, 244 inner ear 258, 267 localization 236 subcellular distribution 237 289

Subject Index EAAT2 (GLT) 49, 219, concentration 236, 244 inner ear 258 localization 233 subcellular distribution 235 EAAT3 (EAAC) 22 localization 238 EAAT4 22 localization 239 subcellular distribution 239 EAAT5 232 localization 240 intersynaptic crosstalk 245 phosphorylation 243 regulation expression 241 posttranslational regulation 242 glutaminase 203 inner ear 258, 260 localization 6, 204 non-neuronal distribution 218 glutamine glutamate precursor 5, 9, 203 immunocytochemistry 276 glutamine cycle 7, 219, 258 glutamine synthetase 219 glutamine transport 5, 258 GRIP 171,191 hair cells 255 hippocampus 47, 54, 119, 125 synapse development 166 Homer 164, 188, 194 homocysteic acid 22 Huntington's disease 25 hypothalamus 24 IP3 receptor 194

kainate receptors 99 cerebellum hippocampus 125 neocortex 118 retina 113 kanamycin 267 ~-ketoglutarate 221 ~-ketoglutarate dehydrogenase 4 290

lactate 3,11 lamprey 273 leucine 5 liprins 192 LTP 53, 109, 169, 186

NR1 splice variants 107 NR2 subunits 109 NR3 110 retina 111 spinal cord 134 NSF 192

MAGUKs 163, 184 MALS 190 metabotropic glutamate receptors 63 anchoring 194 classification 63 mGluR1 localization 76 mGluR2 localization 80 mGluR3 localization 80 mGluR4 localization 83 mGluR5 localization 78 mGluR6 localization 84 mGluR7 localization 84 mGluR8 localization 86 postsynaptic 87 presynaptic 88, 282 second messengers 64 splice variants 65 subcellular distribution 87 target-cell specific segregation 89 methionine sulfoximine 6

receptor anchoring 183 receptor targeting 169, 183 reeler mouse 131 retina 27, 57, 77, 78, 80, 111, 157, 218, 223 Rim 279 RNA editing 111

neurofilament NF-L 190 neuromuscular junction 157, 185 nitric oxide 54 nitric oxide synthase 187 NMDA receptors 99, 104 anchoring 165, 183 cerebellum 130 GABAergic interneurons 121,131 gating 189 hippocampus 119 inner ear 266 mRNA distribution 106 neocortex 114

SAP-97 163, 185 SAP-102 185 Shank 195 silent synapses 162, 166 solitary tract 18 somatostatin 90 spectrin 190 spinal cord 13, 56, 133, 217 Clarke's column 14 intermediolateral nucleus 16 intrinsic neurons 14 primary afferents 13 spinocervical tract 17 S-SCAM 190 suhthalamic nucleus 26

organ of Corti 255 Parkinson's disease 25 PDZ domains 184, 192 pedunculopontine nucleus 27 photoreceptors 28 Piccolo 279 PICK1 192 pinealocytes 49, 76, 152 pontine nucleus 19 presbyacusis 267 PSD-93 185 PSD-95 163, 184, 190 guanylate kinase-like domain 188 knockout 186 Purkinje cells 77, 87, 130, 161 pyruvate carhoxylation 7 in neurons 7

Subject Index synapse development 165 synapsin 279 synaptic vesicles endocytosis 286 recycling 284

SynGAP 187

tinnitus 267

taste buds 157 thalamocortical projection 26 thalamus 23, 152

vestibular end-organ 265

yotiao 190

291

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