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
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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
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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.
Biochemistry and anatomy of transmitter glutamate
Ch. I
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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J. Broman et al.
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
Biochemistry and anatomy of transmitter glutamate
Ch. I
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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J. Broman et al.
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
Ch. I
/
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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J. Broman et al.
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 ~~" .; - ""
\ 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
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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
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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
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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
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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.
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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
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Fig. 1. AMPA receptor subunit mRNA dist
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=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).
102
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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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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
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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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C2
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).
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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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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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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.
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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
o
D~
, rr~'L '
"0
r
~.~o
Regional and synaptic expression of ionotropic glutamate receptors
~
Ch. V
5mm
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
Ch. V
R.S. Petralia et al.
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
Regional and synaptic expression of ionotropic glutamate receptors
Ch. V
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
Ch. V
R.S. Petralia et al. L
~
h
,
,j
.,.. . . .
..~ ~,,.~
.,
~
,~
~
,
,
~i~
~
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
Regional and synaptic expression of ionotropic glutamate receptors
Ch. V
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
~ c~-actinin ~
r
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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PlO
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9 . ' p
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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
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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
Regional and synaptic expression of ionotropic glutamate receptors
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