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I N T E G R A T E D SYSTEMS OF THE CNS PART III

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HANDBOOK OF CHEMICAL NEUROANATOMY Series Editors" A. Bj6rklund and T. H6kfelt

Volume 12

INTEGRATED SYSTEMS OF THE CNS, PART III

Cerebellum, Basal Ganglia, Olfactory System Editors."

L.W. S W A N S O N Department of Biological Sciences, University of Southern California, Los Angeles, CA, U.S.A. oo

A. B J O R K L U N D Department of Medical Cell Research, University of Lund, Lund, Sweden to

T. H O K F E L T Department of Neuroscience, Histology, Karolinska Institute, Stockholm, Sweden

1996

ELSEVIER A m s t e r d a m - Lausanne - New York - O x f o r d - S h a n n o n - Tokyo

9 1996 Elsevier Science B.V. All rights reserved. No part of this publication may be reproduced, stored in a retrieval system or transmitted in any form or by any means, electronic, mechanical, photocopying, recording or otherwise without the prior written permission of the Publisher, Elsevier Science B.V., Copyright and Permissions Department, P.O. Box 521, 1000 AM Amsterdam, The Netherlands. No responsibility is assumed by the Publisher for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions or ideas contained in the material herein. Because of the rapid advances in the medical sciences, the Publisher recommends that independent verification of diagnoses and drug dosages should be made.

Special regulations for readers in the USA. This publication has been registered with the Copyright Clearance Center, Inc. (CCC), 222 Rosewood Drive, Danvers, MA 01923. Information can be obtained from the CCC about conditions under which photocopies of parts of this publication may be made in the USA. All other copyright questions, including photocopying outside the USA, should be referred to the Publisher.

ISBN 0-444-82451-0 (volume) ISBN 0-444-90340-2 (series) This book is printed on acid-free paper.

Published by: Elsevier Science B.V. RO. Box 211 1000 AE Amsterdam The Netherlands

Printed in The Netherlands

Dedicated to J/mos Szentfigothai and Walle J.H. Nauta

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List of contributors MATTHEW ENNIS Department of Anatomy The University of Maryland School of Medicine Baltimore, MD 21201 U.S.A.

MICHAEL T. SHIPLEY Department of Anatomy The University of Maryland School of Medicine Baltimore, MD 21201 U.S.A.

CHARLES R. GERFEN Laboratory of Systems Neuroscience National Institute of Mental Health Bldg 36 Room 2D-10 Bethesda, MD 20892 U.S.A.

J. V O O G D

D. JAARSMA Department of Anatomy Erasmus University Medical Center RO. Box 1738 3000 DR Rotterdam The Netherlands

CHARLES J. WILSON Department of Anatomy and Neurobiology University of Tennessee School of Medicine Memphis, TN U.S.A.

E. MARANI Department of Physiology Leiden University Rijnsburgerweg 10 2300 RC Leiden The Netherlands

Department of Anatomy Erasmus University Medical Center P.O. Box 1738 3000 DR Rotterdam The Netherlands

LEE A. ZIMMER Department of Anatomy The University of Maryland School of Medicine Baltimore, MD 21201 U.S.A.

JOHN H. MCLEAN Division of Basic Medical Sciences Memorial University of Newfoundland St. John's, Newfoundland Canada A 1B 3V6

vii

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Preface It is with a mixture of pleasure and sadness that we dedicate this third volume of the

Integrated Systems series of the Handbook of Chemical Neuroanatomy to the memory of two outstanding structural neuroscientists, J~mos Szentfigothai and Walle J.H. Nauta, who are widely regarded as having led the Romantic School of neuroanatomy through the Twentieth Century. Szentfigothai was born on October 31, 1912, in Budapest, and passed away on September 8, 1994 in his native city. He was a student of Cajal's friend von Lenhoss6k, and like Cajal made enduring contributions to our understanding of many components of the nervous system, including (roughly in chronological order) the autonomic system, spinal cord, vestibulo-ocular and stretch reflex circuitry, neuroendocrine system, cerebellum, thalamus, and cerebral cortex. What sets his work apart from many of his contemporaries was the ability to generalize sensibly. This led, for example, to the concepts of synaptic glomeruli and neuronal modules, and to the synthesis for which he will always be remembered, The Cerebellum as a Neuronal Machine, published in 1967 with his collaborators John Eccles and Masao Ito. Nauta was born on June 8, 1916 in Medan, Indonesia; received the M.D. and Ph.D. degrees at the University of Utrecht; served the last 30 years of his career at the Massachusetts Institute of Technology; and died on March 24, 1994. He perhaps will be remembered longest for the 'Nauta method', the first selective silver impregnation technique for degenerating axons. It was introduced in 1950 and variants were the method of choice for tracing axonal connections for about 25 years, until the use of more sensitive intraaxonal transport techniques became widespread. However, Nauta was a brilliant writer and an inspiring lecturer; and he published very influential experimental analyses of many forebrain systems in a variety of mammals. The limbic system and basal ganglia were his specialties, and indeed his work with Mehler on the lentiform nucleus of the cat and monkey was the first paper published in Brain Research (I :3-42, 1966) and is a classic with regard to both style and content. We are profoundly grateful to the authors who have committed so much time and thoughtfulness to the chapters in the third part of the Integrated Systems component of the Handbook. When planning began in 1983, we had hoped to review each of the major sensory and motor systems, along with parts of the broader system that controls motivated and emotional behavior. Furthermore, each chapter was to be written from a dual perspective- a classical functional neuroanatomical overview, combined with what has been learned more recently about neurotransmitters and receptors within the circuitry. For the usual reasons familiar to editors, all of the planned chapters were not written, and it proved impossible to devote single volumes to an internally consistent theme. Nevertheless, the series as a whole does survey the major sensory systems (retina by Ehinger and Dowling, part I; central visualpathways by Parnavelas, Dinopoulos, and Davies, part II; auditory system by Aitkin, part II; somatosensory system by Rustioni and Weinberg, part II; gustatory and related chemosensory systems by Kruger and Mantyh, part II; and olfactory system by Shipley, McLean, Zimmer, and Ennis, part III); two important parts of the motor system (cerebellum by Voogd, Jaarsma, and Marani, part III; basal ganglia by Gerfen and Wilson, part III); and three key parts of

ix

the limbic system (hypothalamusby Swanson, part I; amygdalaby Price, Russchen, and Amaral, part I; hippocampusby Swanson, K6hler, and Bj6rklund, part I). The literature in the field as a whole continues to explode. Keeping pace is a challenge that we hope will be facilitated by the imminent revolutions in electronic publishing, database management, and computer graphics. Los Angeles, Lund and Stockholm in June 1995 LARRY W. SWANSON

ANDERS BJORKLUND

TOMAS HOKFELT

Contents THE CEREBELLUM, CHEMOARCHITECTURE AND A N A T O M Y J. VOOGD, D. JAARSMA AND E. MARANI ~

2. 3.

Introduction Cytology of the cerebellar cortex Chemical anatomy of the cerebellar cortex 3.1. Purkinje cells 3.1.1. Gamma-aminobutyric acid (GABA), glutamic acid decarboxylase (GAD) and the GABA-transporters in Purkinje cells 3.1.2. Motilin and taurine in Purkinje cells 3.1.3. Calcitonin gene-related peptide (CGRP), acetylcholinesterase (ACHE), somatostatin and tyrosine hydroxylase in Purkinje cells 3.1.4. The localization of the IP3 receptor and the intracellular calcium stores of Purkinje cells 3.1.5. Protein kinase C in Purkinje cells 3.1.6. cGMP, cGMP-dependent protein kinase and nitric oxide synthase in Purkinje cells 3.1.7. Calcium-binding proteins in Purkinje cells 3.1.8. Other specific biochemical markers for Purkinje cells 3.1.9. Cytoskeleton and metabolism of Purkinje cells 3.1.10. Nerve growth factor and nerve growth factor-receptor protein in Purkinje cells 3.1.11. Immunoreactivity of Purkinje cells in paraneoplastic diseases 3.2 Excitatory pathways 3.2.1. Mossy fibers 3.2.2. Climbing fibers 3.2.3. Granule cells and parallel fibers 3.3. Localization of glutamate receptors 3.3.1 Ionotropic glutamate receptors 3.3.2. Metabotropic glutamate receptors 3.4. Nitric oxide: the cerebellar localization of nitric oxide synthase, guanylate cyclase and cyclic GMP 3.5. Adenosine, 5'-nucleotidase and adenosine desaminase 3.6. Interneurons of the cerebellar cortex 3.6.1 Stellate and basket cells 3.6.2. Golgi cells and Lugaro cells 3.6.3. Unipolar brush cells 3.7. Localization of GABA receptors and glycine receptors 3.7.1. GABAA receptors

1 1

17 17

17 21

23 24 32 34 36 38 43 44 47 49 51 55 57 60 60 72 76 77 81 84 85 89 93 93

xi

4. 5.

6.

xii

3.7.2. GABAB receptors 3.7.3. Glycine receptors 3.8. Monoaminergic afferent systems and receptors 3.9. Hypothalamocerebellar connections and histaminergic projections 3.10. Cholinergic systems and acetylcholinesterase (ACHE) in the cerebellum 3.10.1. Distribution of choline acetyltransferase 3.10.2. Cholinergic receptors 3.10.3. Acetylcholinesterase 3.11. Neuroglia Gross anatomy of the mammalian cerebellum The cerebellar nuclei 5.1. Subdivision of the cerebellar nuclei 5.1.1. The cerebellar nuclei of the cat 5.1.2. The cerebellar nuclei of primates 5.1.3. The cerebellar nuclei of the rat 5.2. The GABAergic nucleo-olivary projection neurons of the cerebellar nuclei 5.3. Nucleocortical and intrinsic neurons of the cerebellar nuclei 5.4. Non-GABAergic projection neurons of the cerebellar nuclei 5.5. Afferent connections of the cerebellar nuclei: Purkinje cell axons 5.6. Extracerebellar afferents of the cerebellar nuclei: collaterals of mossy and climbing fibers 5.7. Extracerebellar afferents of the cerebellar nuclei: serotoninergic, noradrenergic, dopaminergic and peptidergic projections Efferent and afferent connections of the cerebellar cortex: corticonuclear, olivocerebellar and mossy fiber connections and cytochemical maps 6.1. Compartments and corticonuclear projection zones: Correlations with cytochemical maps 6.1.1. Corticonuclear projection zones in the cat. Correlation with white matter compartments and cytochemical zones 6.1.2. Compartments and corticonuclear projection zones in monkeys 6.1.3. Parasagittal zonation in the cerebellar cortex: Antigenic compartmentation for Zebrin and other markers 6.1.4. The corticonuclear projection of the cerebellum of the rat. Correlations with Zebrin-antigenic compartmentalization 6.1.5. The corticovestibular and corticonuclear projections of the flocculus and the caudal vermis. Correlations with cytochemical zones and compartments 6.2, Regional differences in the development of the cerebellum 6.3. The organization of the olivocerebellar projection 6.3.1. Configuration and ultrastructure of the inferior olive 6.3.2. Afferent connections of the inferior olive

100 101 102 111 113 113 121 127 128 133 138 140 146 148 151 154 158 160 164

165 167

170 177

177 184 189

201

207 217 225 225 233

6.3.3. 6.3.4. 6.4.

Mossy 6.4.1. 6.4.2. 6.4.3. 6.4.4.

.

,

9.

The connections between the inferior olive and the cerebellum The distribution of peptides and calcium binding proteins in climbing fibers and cells of the inferior olive fiber systems Concentric and discontinuous, lobular arrangement of mossy fiber systems Zonal arrangement in the termination of mossy fibers: Correlations with cytochemical maps The somatotopical organization in mossy fiber pathways Collateral projections of mossy fiber systems to the cerebellar nuclei. The nuclear projection of the red nucleus The chemoarchitecture of mossy fibers

6.4.5. Postscript 7.1. Biochemical correlates of cell types and fiber systems 7.2. Neurotransmitters and their receptors 7.3. Lobules and zones 7.4. The role of biochemically defined systems in cerebellar motor control Acknowledgements References

242 275 284 284 293 299

302 303 305 305 307 307 309 310 311

II. THE BASAL G A N G L I A - C.R. GERFEN AND C.J. WILSON 1. 2. 3. 4.

5.

6.

7.

Introduction Organizational overview 2.1. Comparisons between rodents and primates Cerebral cortex input to striatum Striatum 4.1. Spiny projection neuron 4.1.1. Cortical input 4.1.2. Thalamic input 4.1.3. Nigrostriatal dopamine input 4.1.4. Spiny cell local collaterals inputs (GABA and peptide) 4.1.5. Cholinergic input 4.1.6. Striatal GABA interneuron inputs 4.1.7. Somatostatin interneuron inputs 4.1.8. Other inputs 4.2. Striatal interneurons 4.2.1. Cholinergic neurons Globus pallidus (external segment) 5.1. Synaptic input 5.2. Output Subthalamic nucleus 6.1. Synaptic input 6.2. Output Substantia nigra/entopeduncular nucleus

371 372 376 377 379 380 382 382 386 388 389 389 389 390 390 394 396 397 399 400 400 402 402

xiii

10.

11.

12. 13.

7.1. Synaptic input to pars reticulata neurons 7.2. Synaptic input to pars compacta neurons 7.3. Projections of pars reticulata neurons Connectional organization of basal ganglia Relationship between cortex and basal ganglia 9.1. Topographic organization 9.2. Overlap of inputs: cortico-cortical organization 9.3. Striatal output systems: topography/convergence/divergence 9.4. Striatal outputs in relation to nigral outputs: dual output systems 9.5. Summary of organization of cortico-basal ganglia circuits Striatal patch/matrix compartments 10.1. Nigrostriatal dopamine system 10.2. Striatal outputs 10.3. Cortical inputs 10.4. Thalamic afferents 10.5. General patch-matrix organization 10.6. Cortical organization related to striatal patch-matrix compartments Direct/indirect striatal output systems 11.1. Connectional basis 11.2. Peptide basis 11.3. Dopamine receptor-mediated regulation 11.4. Other (non-dopaminergic) regulatory receptor systems in striatum 11.5. Cellular interactions within the striatum 11.6. Functional significance 11.7. Regional differences Acknowledgements References

403 404 407 409 409 410 413 418 421 425 426 427 429 431 435 435 437 439 439 443 447 449 451 453 455 457 457

III. THE OLFACTORY S Y S T E M - M.T. SHIPLEY, J.H. MCLEAN, L.A. ZIMMER AND M. ENNIS 1.

2.

xiv

469 Introduction 470 1.1. The olfactory epithelium 473 1.2. Two olfactory systems 473 1.3. Human diseases and the olfactory system 474 The main olfactory bulb 474 2.1. Laminar organization 474 2.1.1. Olfactory nerve layer 475 2.1.2. Glomerular layer 486 2.1.3. External plexiform layer 488 2.1.4. Mitral cell layer 490 2.1.5. Internal plexiform layer 491 2.1.6. Granule cell layer 2.1.7. Mitral-granule cell interactions: Anatomical considerations 492 493 2.1.8. Subependymal zone 493 2.2. Transmitter receptors in the MOB 493 2.2.1. Excitatory amino acids (EAAs) 493 2.2.2. GABA receptors

2.3.

Influence of the olfactory nerve on transmitter expression in MOB neurons 2.4. Functional organization of the MOB 2.4.1. Organization of olfactory nerve inputs to MOB 2.4.2. Broad topographic mapping 2.4.3. Neural processing in the glomerular layer 2.4.4. The mitral/granule cell inhibitory system 2.4.5. Glomerular versus infraglomerular inhibition 2.5. Outputs of the MOB 2.5.1. Intrabulbar collaterals 2.5.2. Mitral/tufted cell projections beyond the MOB 2.5.3. Projections to olfactory cortex 2.5.4. Transmitter(s) mediating MOB to PC monosynaptic excitation 2.6. Centrifugal afferents to MOB 3. Primary olfactory cortex 3.1. Anterior olfactory nucleus (AON) 3.1.1. Architecture of AON 3.1.2. Inputs to AON 3.1.3. Outputs of AON 3.1.4. Organization of AON circuitry 3.1.5. Transmitters of AON 3.1.6. Transmitter receptors in AON 3.1.7. Functions of AON 3.2. Rostral olfactory cortex 3.2.1. Indusium griseum 3.2.2. Anterior hippocampal continuation 3.2.3. Taenia tecta 3.2.4. Infralimbic cortex 3.2.5. Olfactory tubercle 3.2.6. Nucleus of the lateral olfactory tract (NLOT) 3.3. Lateral olfactory cortex 3.3.1. Architecture of the lateral olfactory cortex 3.3.2. Neuron types in the piriform cortex 3.3.3. Connections of the lateral olfactory cortex 3.3.4. Transmitter receptors in the lateral olfactory cortex 3.3.5. Piriform cortex is a seizurogenic focus 3.3.6. Modeling of olfactory network function 4. Integration of the main olfactory system with other functions 4.1. Odors and cognition 4.2. Olfaction and taste/visceral integration 4.3. Olfaction and motor activity 4.4. Olfaction and memory 5. The accessory olfactory system 5.1. Accessory olfactory bulb 5.1.1. Neurotransmitters in the AOB 5.1.2. Transmitter receptors in the AOB 5.1.3. Outputs of the AOB 5.1.4. Centrifugal afferents to AOB

493 496 496 496 498 501 503 504 504 504 505 506 507 507 509 509 509 509 510 514 514 515 516 516 516 516 518 518 519 519 519 522 524 529 529 532 532 532 534 534 536 536 536 537 538 539 539

XV

5.2.

,

7. 8. 9.

Higher order connections of the accessory olfactory system and reproductive functions 539 541 5.3. Sexual dimorphism of AOB and its target structures 541 'Non-olfactory' modulatory inputs to the olfactory system 541 6.1. Cholinergic innervation of the olfactory system 541 6.1.1. Cholinergic inputs to the MOB 544 6.1.2. Cholinergic inputs to the piriform cortex 546 6.2. Noradrenergic (NE) innervation of the olfactory system 546 6.2.1. NE innervation of the olfactory bulb 548 6.2.2. NE inputs to the piriform cortex 550 6.3. Serotonin (5-HT) innervation of the olfactory system 6.3.1. 5-HT innervation of the MOB 550 551 6.3.2. 5-HT inputs to the piriform cortex 553 6.4. Dopamine (DA) innervation of the olfactory system 6.4.1. Dopamine (DA) innervation of the piriform cortex 553 Comparison of NE, 5-HT and DA inputs in the rat piriform cortex 553 6.5. 553 6.6. Differential innervation of MOB and AOB 555 Acknowledgments 555 Abbreviations 556 References

SUBJECT INDEX

xvi

575

CHAPTER I

The cerebellum: chemoarchitecture and anatomy J. VOOGD, D. JAARSMA AND E. MARANI

......... but the Spirits inhabiting the Cerebel perform unperceivedly and silently their Work of Nature without our Knowledge or Care. Thomas Willis. Of the Anatomy of the Brain. Englished by Samual Pordage, Esquire, London. Printed for Dring, Harper, Leigh and Martyn, 1681. Facsimile Edition, McGill University Press, Montreal, 1965. p. 111.

1. INTRODUCTION During the last 150 years the morphology of the cerebellum attracted numerous histologists. Its relatively simple structure, with its three-layered cortex and clearly defined afferent and efferent connections made it one of the favourite sites in the brain to test out new hypotheses on the connectivity, the development and chemical interaction in nervous tissue. We have attempted to review present knowledge about the external and internal morphology of the cerebellum and to relate the 'classical' topography of the cerebellum to the more recently discovered chemical specificity of its neurons and afferent and efferent pathways. Not all what is new in the histochemistry of the cerebellum is relevant to a better understanding of its chemoarchitecture. This review, therefore, does not pretend to be complete. It is focussed on afferent and intrinsic connections of the cerebellum. The efferent connections of the cerebellum to the brain stem and the spinal cord have not been systematically covered.

2. CYTOLOGY OF THE CEREBELLAR CORTEX A complete description of the histology of the cerebellar cortex was given by Ramon y Cajal (1911) (Figs 1 and 4). More recently the anatomy of the cortex including its ultrastructure was reviewed by Braitenberg and Atwood (1958), Eccles et al. (1967), Fox et al. (1967), Mugnaini (1972), and Palay and Chan-Palay (1974). Three layers are distinguished in the cortex (Fig. 3). The granular layer borders on the central white matter of the cerebellum. The Purkinje cell layer contains the cell bodies of the Purkinje cells, that are arranged in a single row. The perikarya of the Bergmann glia (the Golgi epithelial cells) are intercallated between the larger Purkinje cells (Fig. 9A). The molecular layer has a low cell content. It contains the dendritic arbors of the Purkinje cells and the Bergmann glial fibers, which run up to the pial surface where they constitute the external glial limiting membrane. The morphology of the cerebellar cortex can be characterized as a lattice: '... it can only be represented in two planes perpendicular to each other and having definite relations to the longitudinal and transversal axes of the

Handbook of Chemical Neuroanatomy, Vo112. Integrated Systems of the CNS, Part IH L.W. Swanson, A. Bj6rklund and T. H6kfelt, editors 9 1996 Elsevier Science B.V. All rights reserved.

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Fig. 1. Cerebellar cortical circuits. Top. Diagram showing the main mossy fiber-granule cell-Purkinje cell circuit and the innervation of the granule cells by the axonal plexus of the Golgi cell. A: mossy fiber; a: granule cell; B: Purkinje cell axon; b: parallel fiber; c: Golgi cell; d: Purkinje cell. Bottom. Similar diagram showing the main cortical circuit and the connection of the basket cell with the Purkinje cell somata. A: mossy fiber; a: granule cell; B: Purkinje cell axon; b: basket cell; C: climbing fiber; c: Purkinje cell soma. Redrawn from Ramon y Cajal (1911).

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Fig. 2. D i a g r a m s of the cerebellar circuit. Inhibitory neurons are indicated in black. A. M a i n circuit. B. Cortical interneurons and recurrent pathways. Abbreviations: B = basket cell; cf = climbing fiber; G = Golgi cell; G R = granule cell; IO = inferior olive; m f = mossy fiber; nc = nucleocortical axons; no = nucleo-olivary axons; pcc = recurrent Purkinje cell axon collaterals; P cell = Purkinje cell; P C N = precerebellar nuclei; p f - parallel fiber; pi = pinceau of basket cell axons; S = stellate cell; U B C = unipolar brush cell; 1 = extracerebellar mossy fiber; 2 - nucleo-cortical mossy fiber; 3 - mossy fiber collateral of uni-polar brush cell.

animal. The whole three dimensional structure, therefore, cannot be obtained by rotation but by translation in two directions, thus producing a lattice' (Braitenberg and Atwood, 1958, p. 1). The elements of the main cerebellar circuit were discovered by Ramon y Cajal (1888, 1911). The electrophysiological properties of the circuit were established by Eccles et al. (1967). The main circuit (Figs 1 and 2) consists of the mossy fiber afferent system, that terminates on the granule cells; the granule cell axons that ascend to the molecular layer and bifurcate into parallel fibers, that run in the long axis of the folium and terminate on the Purkinje cells and the projection of the Purkinje cells to the cerebellar or vestibular nuclei. Each Purkinje cell is innervated by a single climbing fiber (Ramon y Cajal, 1911; Eccles et al., 1966a) that takes its origin from the contralateral inferior olive. The synaptic connections of mossy fibers, parallel fibers and climbing fibers are excitatory. The Purkinje cells are inhibitory and use gamma aminobutyric acid (GABA) as a transmitter (Ito and Yoshida, 1964). Small interneurons of the cerebellar cortex (stellate, basket and Golgi cells) receive a parallel fiber input and constitute inhibitory feed back and feed forward loops terminating on the granule cells and the Purkinje cells (Figs 1, 2 and 4). The main determinant of the firing rate of Purkinje cells is the mossy fiberparallel fiber system. Excitatory coupling between climbing fibers and Purkinje cells is very strong, but the frequency of the complex spikes evoked in Purkinje cells by the climbing fiber is too low to contribute significantly to its firing rate. The function of the climbing fibers, therefore, is one of the main problems in cerebellar neurobiology. Purkinje cells project to the cerebellar or the vestibular nuclei, where their axons terminate with inhibitory synapses. The cerebellar nuclei receive their excitatory drive from collaterals of the mossy and the climbing fibers.

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Fig. 3. A. Nissl-stained section of the cerebellar cortex of the cat. G = Golgi cell; Gr = granule cells; P = Purkinje cell, asterisks: protoplasmatic islands of Held. Bar = 20 r Purkinje (1837).

B. diagram of the cerebeUar cortex of

Granule cells are small neurons located in cell nests in the granular layer. Cell-free spaces in the granular layer, that are known as the protoplasmatic islands of Held, contain the terminals of the mossy fibers (Fig. 3A, asterisks). Mossy fibers originate from many different sites in the spinal cord and the brain stem and constitute the main afferent system of the cerebellar cortex. Mossy fibers are myelinated fibers that branch extensively within the cerebellar white matter and the granular layer. They terminate with large irregular swellings (the mossy fiber rosettes, Figs 1, 5 and 6) that are located along or at the end of the axon. Each rosette forms the center of a complex synapse (cerebellar glomerulus) between the mossy fiber rosette, the dendrites of several granule cells and the terminals of one type of short axon (Golgi) cell of the cerebellar cortex. More than one mossy fiber rosette may be present within a protoplasmatic island. Granule cells possess 3-4 short dendrites, terminating in claw-like excrescenses (Fig.7). The thin, unmyelinated axon ascends towards the molecular layer, where it bifurcates in the form of a T. The two branches, that are known as the parallel fiber, pursue a straight course in the long axis of the folia, parallel to the thousands of other parallel fibers that constitute the bulk of the molecular layer. Parallel fibers synapse with dendrites of Purkinje cells and short axon cells in the molecular layer. Both the ascending portion of the granule cell axon and the parallel fiber are beaded. These varicosities probably correspond to the synaptic sites (Fig. 7D-E). Parallel fibers are very long. In monkeys their length varied between 0.8 and 5 mm. (Fox and Barnard, 1957). Maximal lengths of parallel fibers of 4.6-5.0 mm were reported for the rat (Brand et al., 1976; Schild, 1980; Mugnaini, 1983). The mean length

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from Golgi-stained material 9 A: molecular layer; B: granular layer; C: white matter; a: Purkinje cell; b: basket cells of the lower molecular layer; d: terminal basket formation of the basket cell axon; e: superficial stellate cells; f: Golgi cell; g: granule cells with their ascending axons; h: mossy fibers; i: the bifurcation of the granule cell axons; j: epithelial glial cell; m: astrocyte of the granular layer; n: climbing fiber; o: branching point of Purkinje cell recurrent axon collaterals. Redrawn from Ramon y Cajal (1911).

of parallel fibers of 4.4 mm, measured after microinjections of biocytin in the granular layer in the rat (Pichitpornchai et al., 1994) is close to the mean length of these fibers of 5 mm, estimated with stereological techniques by Harvey and Napper (1988). The two branches of the parallel fiber are of equal length (Pichitpornchai et al., 1994). Shorter parallel fibers are located at the base of the molecular layer (mean branch length 2.08 mm), they become progressively longer as they approach the pial surface (mean branch length 2.35 mm: Pichitpornchai et al., 1994). Parallel fibers in the superficial molecular layer are of a smaller calibre than deep parallel fibers (Fox and Barnard, 1957, monkey). A similar increase in size of the parallel fibers from superficial to deep laminae of the molecular layer was noticed by Pichitpornchai et al. (1994) in the rat. They also observed proximo-distal tapering of parallel fibers. Van der Want et al. (1985a,b) observed corresponding differences in synaptic size in superficial and deep layers of the molecular layer in the cat. The size and the spacing of the varicosities along the parallal fibers was found to be correlated with their caliber. The mean interval between two varicosities was 5.2 ~tm for the parallel fibers, 4.02 ~tm for the ascending axon of the granule cell (Pichitpornchai et al., 1994). The lamination in the molecular layer may be the expression of a deep to superficial gradient in the development of the parallel fibers

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J. Voogd, D. Jaarsma and E. Marani

Fig. 5. Mossy fiber rosettes in the granular layer. Left. Mossy fiber rosettes from neurons of the lateral reticular nucleus, labelled with antegrade transport of Phasaeolus vulgaris lectin. Bar = 25/lm. Right: Mossy fibers, Golgi impregnation. Cajal (1911). Abbreviations: a = large, terminal rosettes; b = rosettes 'en passage'; c = small rosette 'en passage'; G = granular layer; M - molecular layer; W = white matter. Courtesy of Dr. T.J.H. Ruigrok.

(Pellegrino and Altman, 1979). A population of thick, short parallel fibers was noticed by Pitchitpornchai et al. (1994) in the deep parts of the molecular layer. Deep lying parallel fibers may be myelinated and are one of the constituents of the supraganglionic plexus located above the Purkinje cells (Mugnaini, 1972). The mossy fiber-parallel fiber-Purkinje cell pathway is characterized by a large divergence. Each mossy fiber terminates on a great number of granule cells and each granule cell contacts hundreds of Purkinje cells along its parallel fiber. An average parallel fiber with a length of 6 mm forms approximately 1100 boutons (Brand et al., 1976). A portion of the molecular layer 6 mm wide contains approximately 750 Purkinje cell dendritic sheets (Brand and Mugnaini, 1976). This number is somewhat lower than the number of available boutons, when a parallel fiber would synapse once with each Purkinje cell it meets on its way (Brand et al., 1976). It is higher than the estimate of Napper and Harvey (1988b) in the rat that 15% of the boutons on parallel fibers synapse with non-Purkinje cells and that the rest synapses once with half of the Purkinje cell dendritic sheets it meets on its way. The granule cell/Purkinje cell ratio was estimated at 274/1 by Harvey and Napper (1988) and at 350-500/1 for different lobules of rat vermis by Drfige et al. (1986). Napper and Harvey (1988) concluded that there are some 175.000 parallel fiber synapses on a single Purkinje cell of the rat. Fox et al. (1967) arrived at a number of 120.000 in monkeys. The actual strength of the convergence of individual mossy fibers to Purkinje cells depends on the distribution of their mossy fiber rosettes. Electrophysiological studies of Bower and Woolston (1983) in the rat demonstrated that Purkinje cells are most responsive to mossy fiber input that reaches the granule cells located immediately below them. Llinas (1982) explained this strong radial connectivity by the greater number of

The cerebellum." chemoarchitecture and anatomy

Ch.l

A

13

Fig. 6. Drawing of horizontal section through rat cerebellum showing orientation of mossy fibers. A. Elliptical segment or stripe of mossy fiber terminals in the medial portion of the anterior lobe showing the strong caudal-rostral organization of the terminal neuropil. Note the small cluster of granule cell bodies at the open arrow. B. Single mossy fiber from the next adjacent section showing the almost linear caudal-rostral pattern of the related terminals and small groups of parallel fibers (pf). a: View of rat cerebellum from the above showing approximate position of the field illustrated (note square and arrow), b: Medial sagittal section through cerebellum showing approximate location and plane of section. Abbreviations: fp = fissura prima; Isim = lobulus simplex; crI = crus I; fsp = fissura superior posterior; fpl = fissura posterolateralis; pf - parallel fiber. Golgi modification; 21-day-old rat. Scheibel (1977).

synapses with Purkinje cells on the ascending portion of the parallel fiber. However, according to Napper and Harvey (1988) the synapses on ascending portions of parallel fibers would account for only 3% of the total number of synapses of these fibers. Pichitpornchai et al. (1994), who observed a closer spacing of varicosities on the ascending axon and the proximal branches of the parallel fibers than on their distal branches, concluded that parallel fibers will exert a graded synaptic influence on their target Purkinje cells, with the most powerful influence occurring on cells located around the proximal regions of the fibers where they bifurcate. Mossy fiber terminal branches in the granular layer are oriented longitudinally, in the same plane as the Purkinje cells (Scheibel, 1977), (Fig. 6) (see also Section 6.4.2.). Mossy fibers, therefore, preferentially activate longitudinally oriented patches of Purkinje cells. Different types of mossy fiber rosettes were described by Brodal and Drablos (1963) with the Glees and Rheumont-Lhermitte silver impregnations and the Golgi method in rat and cat. Highly branching mossy fibers terminating in small, relatively simple rosettes, located along or at the end of the fiber, occur in all parts of the cerebellum. Large rosettes, consisting of aggregations of larger and smaller argyrophilic particles, interconnected by fiber fragments occur exclusively in nodulus and adjoining uvula, lingula and flocculus. The dendritic tree of the Purkinje cell is flattened in a plane perpendicular to the long

Ch.I

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The cerebellum." chemoarchitecture and anatomy

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Fig. 7. Granule cells and parallel fibers after an injection of biocytin in lobule X of the cerebellum of the rat. A. Biocytin labelled granule cell. B. Golgi-impregnated granule cells and parallel fibers in transverse section. Cajal (1911). C. biocytin injection site in granular layer and labelled parallel fibers in molecular layer. D. bifurcation site of labelled parallel fibers. E. labelled varicose parallel fibers. Abbreviations: A: molecular layer; B: granular layer; C: white matter; a: granule cell axon; b: bifurcation of granule cell axon; d: Purkinje cell; f: Purkinje cell axon; g: granular layer; I: injection site; m: molecular layer. Bars in A = 12/~m, in C = 500 /~m, in D and E - 50/~m. Courtesy of Dr. T.J.H. Ruigrok. (

axis of the folia (Figs 8 and 9). The soma and the proximal dendrites of the Purkinje cell are relatively smooth, the distal dendrites (spiny branchlets) bear long-necked spines (Fox and Barnard, 1957). When the parallel fibers traverse the Purkinje cells they terminate with boutons en passage on the spines of their spiny branchlets. Climbing fibers terminate on short, stubby spines on the proximal dendrites of the Purkinje cells (Larramendi and Victor, 1967; Palay and Chan-Palay, 1974) (Figs 10, 11 and 14). The axon of the Purkinje cell is myelinated (Fig. 9) and gives rise to recurrent collaterals (Bishop, 1982, 1988; Bishop and O'Donoghue, 1986; Bishop et al., 1987; O'Donoghue and Bishop, 1990). The collaterals form a plexus of beaded axons, mainly at the level of the Purkinje cell layer (Fig. 8a and b). They terminate on neighbouring 9

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J. Voogd, D. Jaarsma and E. Marani

Fig. 9. Purkinje cell in sagittal section. H~iggqvist stain. A. The small, densely stained nuclei in the Purkinje cell layer belong to the Bergmann glial cells. B. Initial segment of Purkinje cell myelinated axon (A) surrounded by pinceau of terminal basket cell axons. Abbreviations: A = Purkinje cell axon; B = Bergmann glial fiber; D = Purkinje cell dendrite. Bar - 25 pm.

10

The cerebellum: chemoarchitecture and anatomy

Ch. I

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Fig. 10. Synapses on mouse Purkinje cell. Climbing fibers terminate on short, stubby spines of proximal dendrites (Ds), parallel fibers terminate on spiny branchlets (Bs). Stellate cell and basket cell axons terminate on proximal dendrites and soma. Larramendi and Victor (1967).

Purkinje cells. The recurrent collaterals extend into the molecular layer where they contact basket cells (O'Donoghue et al., 1989). The whole collateral arborization is oriented perpendicular to the long axis of the folia, i.e. in the same plane as the dendritic tree of the Purkinje cell. In the cat it measures 300-700 #m in the sagittal and 100-400 #m in the transverse direction (Bishop, 1988). The width of the arborization and its penetration in the molecular and granular layers varies for different parts of the cerebellum. Recurrent collaterals of Purkinje cell axons are constituents of the infra- and supraganglionic plexus. The main Purkinje cell axon enters and traverses the white matter to terminate on cells of the cerebellar or the vestibular nuclei. Climbing fibers (Fig. 14) innervate the Purkinje cells, each Purkinje cell receiving only one climbing fiber (Ramon y Cajal, 1911). The olivocerebellar parent fibers of the climbing fibers branch extensively in the cerebellar white matter. For the adult rat the ratio of climbing fiber innervated Purkinje cells to neurons of the inferior olive is approximately 10:1 (Schild, 1970; Delhaye-Bouchaud et al., 1985). During their development the Purkinje cells receive more than one climbing fiber, it is not known how these supernumary climbing fibers are eliminated. Branching of olivocerebellar fibers occurs 11

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J. Voogd, D. Jaarsma and E. Marani

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Fig. 11. Diagram of the interaction of Purkinje cell dendrite with a climbing fiber and several parallel fibers. A proximal Purkinje cell dendrite (pd) shows stubby thorns contacted by a climbing fiber (cf), whereas parallel fibers (pf) synapse on spines protruding from a spiny branchlet (sb). Rossi et al. (1991).

in the parasagittal plane (Armstrong et al., 1973; Brodal et al., 1980; Rosina and Provini, 1983). This is one of the reasons for the longitudinal, strip-like organization of the olivocerebellar projection (see Section 6.3.3.). Transverse branching is limited to climbing fibers terminating in certain longitudinal strips (Ekerot and Larson, 1982). Climbing fibers take their origin from the contralateral inferior olive. For a long time the origin of the climbing fibers remained obscure. Their ultrastructure and their mode of termination were first recognized by Larramendi and Victor (1967) (Figs 10 and 11) in the mouse as beaded fibers, with boutons en-passage, filled with rounded vesicles terminating on short spines on Purkinje cell proximal dendrites. The clear intervesicular axoplasm distinguishes climbing fibers from the neurofilamentous basket cell axons. Earlier Scheibel and Scheibel (1954) had reviewed Ramon y Cajal's (1888) original description of the morphology of the climbing fiber. They concluded that climbing fibers emit collaterals in the granular and molecular layer, that terminate in glomeruli, on somata of Golgi, basket and stellate cells and on neighbouring Purkinje cells. Szentagothai and Rajkovits (1959) subsequently identified climbing fibers in axonal degeneration studies from their 'Scheibel-collaterals' and concluded that the climbing fibers originate from the inferior olive. Hamori and Szentagothai (1966b) described the climbing fibers as packed with neurofilaments and making synaptic contacts with few vesicles on the smooth parts of the dendrites. They probably mistook ascending collaterals of basket cell axons for the climbing fibers. The origin of the climbing fibers from the inferior olive was finally settled by Desclin (1974), who observed their degeneration with axonal silver impregnation methods after lesioning the inferior olive of the rat with 3-acetylpyridin (3-AP) administrated intra-peritoneally. In an exhaustive analysis of the normal light- and ultrastructural morphology of the climbing fiber, Palay and Chan-Palay (1974) observed the existence of climbing fiber glomeruli and synapses with Golgi cells in the granular layer and synaptic contacts of climbing fiber tendrils with basket and stellate cells. Desclin and Colin (1980) were unable to confirm these types of collateral contacts, outside the Purkinje cells, in an 12

The cerebellum." chemoarchitecture and anatomy

Ch. I

//

Fig. 12. Purkinje cells from the cerebellum of, from left to right, birds (Gallus domesticus, Feirabend, 1983); mammals (cat, Cajal, 1911) and fish (Gnathonemus petersii, Nieuwenhuys, 1969). Note different length, orientation and position in the molecular layer of the spiny dendritic branchlets.

ultrastructural study of the cerebellar cortex of 3-AP-treated rats. O'Donoghue et al. (1989) found intracellularly stained basket cells of the cat to lack climbing or mossy fiber terminals on their somata. During postnatal maturation of the cerebellum of the mouse, Mason and Gregory (1984) found many axons that combine the morphology and synaptic connections of both climbing and mossy fibers. These combination fibers are rare in the adult. Purkinje cell dendritic trees in the molecular layer remain oriented perpendicular to the parallel fibers irrespective of the changes in direction of the folial chain. Their dendrites share this orientation with the climbing fibers terminating on them. This type of spatial organization is found in all vertebrates and is the main condition which determines the morphology of the cerebellum. Purkinje cells in fish and amphibians are not arranged in a monolayer, but can be clustered in specific parts of the cortex, reminiscent of the clustering of the Purkinje cells during early stages of cerebellar development in all vertebrates (Nieuwenhuys, 1967). Purkinje cells in lower vertebrates differ from the mammalian type by the disposition of their smooth, proximal branches and their spine-loaden terminal branches in the molecular layer (Fig. 12). In fish the proximal smooth branches are found at the same level as the somata of the Purkinje cells, and the distal spiny branchlets extend as straight spikes into the molecular layer. This condition was extensively studied by Nieuwenhuys and Nicholson in the cerebellum of mormyrid fish (Nieuwenhuys and Nicholson, 1969a,b). As a consequence the climbing fibers, that synapse with the smooth proximal part of the dendrites, do not 'climb' 13

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into dendrites within the molecular layer, but terminate at the same level as the perikarya of the Purkinje cells (Kaiserman-Abramof and Palay, 1969). In reptiles and birds the smooth, proximal dendrites with their climbing fiber terminals do not extend beyond the lower third of the molecular layer (Mugnaini, 1972; Freedman et al., 1977; Kfinzle 1985). Only in mammals the smooth branches and the climbing fiber arborizations reach the pial surface (Fig. 12). Cerebellar nuclei that contain the target cells of the Purkinje cell axons have been described in species of all vertebrate classes. In some species of fish target cells of the Purkinje cell axons are also located within the cortex among the Purkinje cells (the 'eurodendroid' cells of Nieuwenhuys et al., 1974). The cells of the 'fourth cortical layer' in some aquatic mammals, that are located below the granule cells in the white matter, can be considered as displaced cerebellar nuclear cells (Ogawa, 1934). Interneurons in the cerebellar cortex are inhibitory and constitute various feed-back and feed-forward circuits between parallel fibers, granule cells and Purkinje cells (Figs 1, 2 and 4). Their dendrites are located in the molecular layer, where they are contacted by parallel fibers. Golgi cells are most numerous in the upper part of the granular layer. Some of their dendrites ramify in the granular layer, where they are contacted by mossy fiber terminals in the glomeruli. The dendritic tree of Golgi cells is not oriented in a specific plane. Recently it was shown by De Zeeuw et al. (1994c) that axons of Golgi cells course for some distance in the supra- or infraganglionic plexus in the direction of the long axis of the folia, before they branch into a dense telodendrion in the granular layer. Their terminals are located at the periphery of the glomeruli, where they synapse with granule cell dendrites (Fox et al., 1967). Their ratio was estimated in the rat as 4-6 Golgi cells for each Purkinje cell. The number of Golgi cells is about three times higher in lobule X than in other lobules (Drfige et al., 1986). However, unipolar brush cells (see below and Section 3.6.3.) may have been mistaken for Golgi cells by these authors.

Fig. 13. Orthogonal arrangement of basket cell axons (thick horizonal fibers oriented in the plane of the Purkinje cells in A and B) and parallel fibers (thin, vertical fibers in A and B). A. Drawing from Golgiimpregnated section, Cajal (1911). B. Bodian-stained section of rat cerebellar cortex. Abbreviations: A and B = stellate cells; C = basket cell axon; E = pericellular basket; F = Purkinje cell dendritic tree; G = climbing fiber; Pb = pericellular baskets. Bar = 100/~m.

14

The cerebellum." chemoarchitecture and anatomy

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Fig. 14. Phaseolus vulgaris lectin-labelled climbing fibers of rat cerebellum. A. Sagittal section. B. Coronal section. Abbreviations: G = granular layer; M = molecular layer; P = Zebrin- labelled Purkinje cells. Bar = 100/lm. Courtesy of Dr. T.J.H. Ruigrok.

Apart from the parallel fiber boutons on the dendrites, the soma of Golgi cells is contacted by Purkinje cell recurrent collaterals (Hamori and Szentagothai, 1966a, 1968, Palay and Chan Palay, 1974). Mossy- and climbing fiber terminals on Golgi cells, that were mentioned by several authors (Hamori and Szentagothai, 1966a, 1968, Palay and Chan Palay, 1974) have not yet been confirmed in experimental axonal tracing studies. Myelinated fibers, indicated as mossy and climbing fibers, and recurrent collaterals of Purkinje cell axons, terminate on Golgi cell somata with large, crenelated synapses ('synapse en marron': Palay and Chan-Palay, 1974). The synapse en marron recently was identified by Mugnaini and Floris (1994) as a synapse of the mossy fibers with the unipolar brush cells of the cerebellar cortex. Stellate cells are located in the entire molecular layer, basket cells constitute a special population located in its lower one third. Dendrites of stellate and basket cells are oriented in a direction perpendicular to the long axis of the folium. Axons of stellate cells terminate on Purkinje cell dendrites. The basket cell axon increases in thickness after it emerges from its soma (Figs 1 and 13). It runs, perpendicular to the long axis of the folium, above the perikarya of the Purkinje cells and gives off descending and ascending collaterals. The descending collaterals branch and surround and synapse with the somata of Purkinje cells. The axons of these pericellular baskets of the Purkinje cell terminate in a periaxonal plexus (the pinceau) surrounding the initial segment of the Purkinje cell axon. Ascending collaterals of the basket cell axon terminate on the smooth surface of the proximal dendrites of Purkinje cells. O'Donoghue et al. (1989) who studied the connections of intracellularly stained basket cells and Purkinje cells in the cat concluded that each basket cell soma received input from recurrent collaterals from a single Purkinje cell. Other afferents of the basket cell include parallel fibers, climbing fibers and stellate and basket cell axons (Palay and Chan Palay, 1974). The infra- and supraganglionic plexus, are located on either side of the layer of Purkinje cell somata. They contain myelinated Purkinje cell collaterals. Most myelinated fibers in the supraganglionic layer are oriented in the long axis of the folia and, therefore, represent myelinated granule cell axons or, possibly, axons of candelabrum cells (Lain6 and Axelrad, 1994) or Golgi cells (De Zeeuw et al. 1994c). In silver impregnations these axons are distinctly smaller than the basket cell axons, that cross them at right angles 15

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(Fig. 13). The so-called multi-layer fibers also contribute to the plexus surrounding the Purkinje cells. These fibers were traced from several sources, including the noradrenergic, serotoninergic and cholinergic cell groups of the brain stem. They ramify in all layers of the cortex, but constitute the densest plexus at the level of the Purkinje cells. They are discussed more fully in Sections 3.8., 3.9. and 3.10.1. Several other neuronal cell types have been identified in the cerebellar cortex. The Lugaro cell is a relatively rare fusiform neuron, located just below the Purkinje cell layer (Lugaro, 1894; Fox, 1959; Palay and Chan-Palay, 1974). Its dendrites stretch out along the boundary of the granular and the Purkinje cell layer, the destination of its axon is not known. Lugaro cells can be discriminated from Golgi cells immunocytochemically with specific antibodies (Section 3.6.2., Fig. 67). The candelabrum cell has been recognized in Golgi-impregnated sections from rat cerebellum by Lain6 and Axelrad (1994). The neuron is rather frequently encountered in all lobules of the cerebellum. Its medium sized perikaryon is sqeezed in between the somata of Purkinje cells. Its dendritic tree is somewhat flattened and mainly extends in the parasagittal plane. One or two dendrites course through the molecular layer, dividing into few, slightly oblique branches, that are covered with irregularly distributed spines. A few slender dendrites branch in the upper granular layer. The axon courses in the direction of the long axis of the folium in the Purkinje ceil - or the supraganglionic layer, and gives off terminal, beaded branches that ascend in the molecular layer at regular parasagittal intervals. The chemical anatomy of the candelabrum cell has not yet been studied (Section 3.6.2). The unipolar brush cells were first identified in the rat by Altman and Bayer (1977) as the 'pale cells' of the granular layer. These cells are intermediate in size between the granule and the Golgi cells, and possess a typical, pale nucleus. They are concentrated in the nodulus, the ventral uvula, the flocculus and parts of the paraflocculus. They are born after the Purkinje cells, but before the stellate, basket and granule cells. The cells were sporadically recognized as monodendritic neurons in a number of immunocytochemical studies (see Section 3.6.3), but have been characterized with Golgi impregnation and electron microscopic methods only recently (Floris et al., 1994; Mugnaini and Floris, 1994; Mugnaini et al., 1994). The name 'unipolar brush cell' was given by Mugnaini and colleagues (Mugnaini and Floris, 1994) after the tip of the stubby dendrite, that forms a tightly packed group of branchlets resembling a paint brush (Fig. 68). The soma of unipolar brush cells is spherical to oval and carries thin appendages. The axon only can be impregnated for a short distance, suggesting that its distal, unimpregnated part is myelinated. Side branches of the axon terminate in rosette-like formations in the granular layer (Fig. 2) (Berthi6 and Axelrad, 1994; Floris et al., 1994; Rossi et al., 1995), the main stem of the axon may enter the white matter. Unipolar brush cells are innervated by one or two mossy fiber rosettes, in the form of particularly extensive contacts. Mossy fibers end on the perikaryon as well as on the dendritic brush (Mugnaini et al., 1994). These large synapses correspond to the 'synapse en marron' of Palay and Chan-Palay (1974), originally identified as a mossy fiber-Golgi cell synapse (see also Monteiro et al., 1986). Unipolar brush cells also receive symmetrical synapses from boutons containing pleomorphic vesicles, presumably originating from Golgi cells or Purkinje cell recurrent axons. Some of the dendritic branchlets may be presynaptic to dendrites of other cells in the granular layer (Floris et al., 1994). Pale cells, monodendritic and unipolar brush cells are all more frequent in the vestibulocerebellum. The chemical identity of the unipolar brush cell will be discussed in Section 3.6.3.

16

The cerebellum." chemoarchitecture and anatomy

Ch. I

3. CHEMICAL ANATOMY OF THE CEREBELLAR CORTEX By virtue of its laminated and relatively simple structure the cerebellar cortex has served as the playground for every student who wanted to test a histochemical reaction or a new antibody on the brain. From this large body of data we have selected those which are important for the understanding of the morphology and the connections of the cerebellum. The localization in the cerebellar cortex of neurotransmitters, peptides, second-messenger systems, calcium-binding proteins and other biochemical markers is reviewed separately for each cell type of the cortex and for the mossy and climbing fibers. Glutamate and GABA receptors, nitric oxide, adenosine, the monoamine afferent systems and receptors, the hypothalamo cerebellar and histaminergic afferents and the cholinergic systems and acetylcholinesterase are discussed in separate sections. The chemoarchitecture of the cerebellar cortex has been reviewed by Schulman (1983), Nieuwenhuys (1985) and Oertel (1993). 3.1. PURKINJE CELLS

3.1.1. Gamma-aminobutyric acid (GABA), glutamic acid decarboxylase (GAD) and the GABA-transporters in Purkinje cells Purkinje cells use gamma-aminobutyric acid (GABA) as their main neurotransmitter and exert a postsynaptic inhibitory effect on cells of the cerebellar and vestibular nuclei (Ito and Yoshida, 1964; Obata et al., 1967; Obata, 1969, 1976; Obata and Takeda, 1969). GABA in rabbit Purkinje cells was first demonstrated using a histochemical method, demonstrating the conversion of GABA into succinic acid (Van Gelder, 1965). In selective uptake studies of [3H]GABA in cerebellar slices, only a low activity was present over the Purkinje cells (H6kfelt and Ljungdahl, 1970, 1971; Schon and Iversen, 1972). Minimal uptake of [3H]GABA was also observed for Purkinje cell axon terminals (Storm-Mathisen, 1975). All Purkinje cell somata of the cerebellum of the rat and their primary and secondary dendrites were immunoreactive for antisera against glutamic acid decarboxylase (GAD), the synthesizing enzyme of GABA (Fig. 62D,E). Varicose fibers and terminals in the cerebellar nuclei were densely stained (Saito et al., 1974; McLaughlin et al., 1974; Oertel et al., 1981b; Perez de la Mora et al., 1981; Somogyi et al., 1985). Immunoreactivity in Purkinje cell somata was generally found to be weak, or to be dependent on blocking of axonal transport by colchicine (Ribak et al., 1978). Strong immunoreactivity in Purkinje cell somata was, however, reported by Mugnaini and Oertel (1985) with an anti-GAD antiserum produced by Oertel et al. (1981 a). The presence of GAD mRNA in Purkinje cells has been demonstrated with in situ hybridization histochemistry in rodents and primates resulting in dense labelling over somata of Purkinje cells (Wuenschell et al., 1986; Julien et al., 1987; Ferraguti et al., 1990; Herrero et al., 1993). Two forms of GAD with apparent molecular weights in the range of 59-67 kDa, that differ by 2-4 kDa, were identified by Chang and Gottlieb (1988) and Martin et al. (1991). In situ hybridization histochemistry with probes for the high molecular weight form, GAD67, and the low molecular weight form, GAD65, showed a prevalent localization of GAD67 over GAD65 in Purkinje cell bodies of rat cerebellum. The reverse localization was reported for Golgi cells (Esclapez et al., 1993; Feldblum et al., 1993). A differential distribution of GAD67 and GAD65 in Purkinje cells was found in immunocytochemical studies with specific antibodies for GAD67 and GAD65. Antibody K2, which is specifc 17

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for GAD67 , strongly immunoreacted with Purkinje cell perikarya, their proximal dendrites and their axon terminals in rat cerebellum (Kaufman et al., 1991; Moffett et al., 1994). The monoclonal antibody GAD-6, which is specific for GAD65 (Chang and 18

The cerebellum." chemoarchitecture and anatomy

Ch. I

Fig. 15. Localization of GABA-like immunoreactivity in semithin sagittal sections of rat (A,B) and chick (C,D) cerebellum. A. Low magnification of cell bodies and neuronal processes reacting with monoclonal anti-GABA antibodies. B. Higher magnification of an area partially included in the frame in A. Note strong immunoreactivity in stellate cell bodies (open arrow), in Golgi neurons (thick arrow), in the basket terminals surrounding Purkinje cell bodies, in puncta at glomeruli (dotted line) and in axons in the white matter. Immunoreactivity in Purkinje cells is weak. C and D. Two typical patterns of GABA-like immunoreactivity observed in Vibratome slices of the chick cerebellum. C: Intensely (thick arrow) and weakly (open arrow) immunoreactive Purkinje cells together with the staining in their dendritic arborization (thick arrowhead). D. Basket terminals around two weakly stained Purkinje cells (open arrowheads). Molecular layer (MO); Purkinje cell layer (P); granular cell layer (GL); white matter (WM). Bar in A = 100 r bar in B, C and D = 25 r (Matute and Streit, (1986). (

Gottlieb, 1988), i m m u n o r e a c t e d with axon terminals of Purkinje cells, but p o o r l y imm u n o s t a i n e d the p e r i k a r y a of Purkinje cells ( K a u f m a n et al., 1991). Antibodies against conjugates of G A B A were first applied to d e m o n s t r a t e specific G A B A - l i k e i m m u n o r e a c t i v i t y in Purkinje cells by S t o r m - M a t h i s e n et al. (1983). Imm u n o r e a c t i v i t y of the cell b o d y and the dendrites with antibodies against conjugates of G A B A was generally f o u n d to be weak or absent, but strong in the a x o n and the myelinated axon collaterals in the infraganglionic, but especially in the supra-ganglionic plexus, and in their terminals in rat (Ottersen and S t o r m - M a t h i s e n , 1984a,b, 1987; Ottersen et al., 1987; M a d s e n et al., 1985; Sdgudla et al., 1985; G a b b o t t et al., 1986; M a t u t e and Streit, 1986; S o m o g y i et al., 1986; A o k i et al., 1986) cat (Somogyi et al., 1985) and m o u s e ( T a k a y a m a , 1994). Staining in Purkinje cell s o m a t a in the chicken was stronger t h a n in m a m m a l s ( M a t u t e and Streit, 1986) (Figs 15, 62, 63). Several G A B A t r a n s p o r t e r proteins, that are active in the high-affinity u p t a k e of GABA

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Fig. 16. Schematic diagram to illustrate the concept of dynamic interrelationships between taurine, motilin, and gamma-aminobutyric acid (GABA) in a single neuron. A neuron with both substances in coexistence may have fluctuating levels of one or both substances depending upon parameters of rhythm, time, and physiologcal demands for one or another mediator during specific types or phases of activity. Chan-Palay (1984). 19

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J. Voogd, D. Jaarsma and E. Marani

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The cerebellum." chemoarchitecture and anatomy

Ch. I

Fig. 17. Immunoreactive staining with taurine (Tau2) antibody in rat cerebellum fixed with 4% paraformaldehyde. A. Tau2-immunoreactive staining on a coronal section though lobule 6 of the vermis. Purkinje cells and dendrites exhibiting Tau-LI were separated by bands of unstained Purkinje cells. M, molecular layer; P, Purkinje cell layer; G, granule cell layer; W, white matter; P", Purkinje cell layer out of plane focus. Location, bregma = 11,5 mm. B. High-power photomicrograph of area indicated in A demonstrating taurine-like immunoreactivity in Purkinje cells (short solid arrows) and dendrites (long solid arrows) separated by unstained Purkinje cells (open arrows). C. Tau2-immunoreactive staining in a horizontal section through lobule 3 of the vermis. D. Adjacent section to (C) demonstrating that taurine-like immunoreactivity was completely absorbed by incubation of Tau2 (1 : 40) with original antigen, taurine conjugated to KLH using glutaraldehydeborohydride. Bars in A, C, and I) = 100/.tm, in B = 50 r Magnusson et al. (1988).

GABA, have been cloned (GAT1-4: Guastella et al., 1990; Lopez-Corcuera et al., 1992; Borden et al., 1992; Liu et al., 1993; and GAT-B: Clark et al., 1992). All transporters occur in brain tissue. The regional distribution of GAT1 was studied by Rattray and Priestly (1993) with in situ hybridization in rat cerebellum. GAT1 mRNA is not expressed by Purkinje cells, but strongly by Bergmann glial cells. GAT-2 may be confined to glia (Liu et al., 1993), but detailed studies of their localization have not been published thus far. 3.1.2. Motilin and taurine in Purkinje cells

Certain inconsistencies in the results on the localization of GABA in Purkinje cells were discussed by Chan-Palay (1984). She concluded that GABA is present in varying amounts in different Purkinje cells and that it may co-exist with other neuroactive substances, notably with motilin and taurine, that also produce an inhibitory action on postsynaptic cells (Fig. 16). The presence of motilin in Purkinje cells was demonstrated with an antibody directed against conjugates of motilin (Chan-Palay et al., 1981; Nilaver et al., 1982). More than half of the Purkinje cells of the rat are immunoreactive for this antibody and in human cerebellum their proportion was even higher (Nilaver et al., 1982). Chan-Palay et al. (1981) found coexistence of GAD and motilin in 10-20% of the Purkinje cells of the rat. The presence of motilin in Purkinje cells has, however, been disputed by Lange (1986), who was unable to demonstrate the presence of motilin using radioimmuno-assay and reversed phase HPLC in extracts of rat cerebellum. Only one of Lange's anti-motilin antibodies, all of which had been demonstrated to be effective in demonstrating motilin-like immunoreactivity in rat duodenum, was found to immunoreact with Purkinje cells in immunocytochemical studies with rat cerebellum. Taurine has been proposed as a neurotransmitter in certain fiber systems. In the guinea pig cerebellum it was found to exert a hyperpolarising effect on Purkinje cell dendrites and was proposed as a neurotransmitter in stellate cell-Purkinje cell synapses (Okamoto et al., 1983). [3H]Taurine was found to accumulate in Purkinje cells. Immunocytochemical studies with antibodies specific for cysteine-sulfonic acid decarboxylase (CSADCase), the enzyme involved in taurine synthesis, by Chan-Palay et al. (1982a,b), showed that CSADCase immunoreactivity was present in most, but not all the Purkinje cells of rat cerebellum, and was more prominent in the main dendritic arbor than in the perikarya and the axon. CSADCase, motilin and GAD-like immunoreactivities were found to co-exist in Purkinje cells located near the midline. In contrast to the observations of Chan-Palay et al. (1982a,b), Almarghini et al. (1991) found CSADCase immunoreactivity to be localized in Bergmann glia and interfascicular oligodendrocytes and to be absent from Purkinje and stellate cells. Most authors who used antisera against conjugates of taurine to localize taurine-like 21

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immunoreactivity found staining in all Purkinje cells of the cerebellum of the rat (Madsen et al., 1985; Campistron et al., 1986b; Tomida and Kimura, 1987; Ida et al., 1987; Ottersen et al., 1988b; Ottersen, 1988, 1989). Magnusson et al. (1988) recognized a zonal distribution of taurine-immunoreactivity similar to the zonal labelling of CSADCase immunoreactivity observed by Chan-Palay et al. (1982a,b) (Fig. 130) in paraformaldehyde-fixed brain tissue (Fig. 17), but not in glutaraldehyde-fixed tissue. Analysis of semithin sections and immunogold electron microscopy indicated that taurine-immunoreactivity is selectively enriched in the somata, proximal and distal dendrites and axon terminals of the Purkinje cells (Fig. 18). Stellate and basket cell somata and their axon terminals are only weakly immunolabelled (Ottersen et al., 1988b; Ottersen 1988, 22

The cerebellum." chemoarchitecture and anatomy

Ch.I

Fig. 18. Photomicrographs showing the distribution of taurine-like immunoreactivity in the rat cerebellum, and the results of different control experiments. A,B. Semithin (0.5 r sagittal sections through vermis posterior treated with taurine (Tau) antiserum 20 diluted 1:3000 and subsequently processed according to the peroxidase-antiperoxidase procedure. A. Intense labelling of the somata, dendrites (small arrowheads), and axons (crossed arrow) of the Purkinje cells. The neurons (large arrowheads) and glial processes (small arrows) of the molecular layer appear immunonegative. Small asterisks indicate pial surface, large asterisk indicates Purkinje cell enlarged in B. B. Note staining of Purkinje cell dendritic spines (large arrow heads). Inset: Semithin test section mounted on the same slide as the tissue section shown in A and B and incubated in the same drops of sera. The test section contains brain protein-glutaraldehyde conjugates of different amino acids, separated by brain tissue that appears as darkly stained zones. Code: 1, GABA; 2, glutamate; 3, taurine; 4, glycine; 5, none (i.e., no amino acid in the reaction mixture); 6, aspartate; 7, glutamine (only small part of section represented in this particular view). The taurine conjugate is selectively stained. C. Transverse semithin section through nucleus interpositus anterior showing intense staining of axons (arrows) and nerve terminal-like dots, some of which (large arrowheads) appear to contact unstained cell bodies (asterisks). Same procedure as in A. D. Thin layer chromatograms (5 mm width) of soluble brain extracts fixed with glutaraldehyde and subsequently stained with an antiserum against glutamate (left strip) or taurine (right strip). The taurine antiserum reveals a single spot which has comigrated with free taurine and which is separate from the spot labelled by the glutamate antiserum. E, T and G indicate the application sites of the brain extract, taurine and glutamate, respectively. E. Adjacent section to that shown in A and accompanying test section (inset) incubated with the taurine antiserum after preabsorption with glutaraldehyde-taurine complexes (200/tM with respect to taurine). There is virtually no staining. Abbreviations: MO, molecular layer; GC, granule cell layer. Bars 25/~m. Ottersen (1988). (

1989). Following hypo-osmotic stress there is a transient shift of taurine from the Purkinje cells to the B e r g m a n n glial c o m p a r t m e n t , where taurine-immunoreactivity now becomes a p p a r e n t (Nagelhus et al., 1993).

3.1.3. Calcitonin gene-related peptide (CGRP), acetylcholinesterase (ACHE), somatostatin and tyrosine hydroxylase in Purkinje cells Some neuroactive substances have been reported to be present in Purkinje cells only for a certain period during development. Calcitonin gene-related peptide ( C G R P ) is almost undetectable in adult rat cerebellum, but a transient immunoreactivity in i m m a t u r e Purkinje cells in rat cerebellum has been detected with antisera against C G R P ( K u b o t o et al., 1987, 1988; Chedotal and Sotelo, 1992). In adult rats, however, C G R P - l i k e immunoreactivity, co-localized with G A D immunoreactivity, can be detected in m a n y Purkinje cells near injections of colchicine (Kawai et al., 1985, 1987). C G R P receptors measured autoradiographically with [125I]CGRP as the ligand, are a b u n d a n t in adult rat and h u m a n cerebellum. [125I]CGRP-binding is dense over the molecular and Purkinje cell layers and low over the granular layer and the cerebellar nuclei (Inagaki et al., 1986). Binding to the molecular layer occurs in a pattern of longitudinal stripes (Kruger et al., 1988) and increases after intraperitoneal administration of harmalin (Rosina et al., 1990, 1992). Choline acetyltransferase (CHAT) and acetylcholinesterase (ACHE) have been found to be transiently expressed in Purkinje cells during development: Purkinje cells in certain parts of the i m m a t u r e rat and guinea pig cerebellum, including the lobules IX and X of the caudal vermis, display a transient reactivity for A C h E (Csillik et al., 1963, 1964; A l t m a n and Das, 1970; Odutola, 1970; Brown et al., 1986). The authors suggested that this transient A C h E activity in Purkinje cells is due to a transient cholinoceptive stage, when they are contacted by cholinergic mossy fiber afferents. A similar, transient expression of C h A T was observed in Purkinje cells of the rat vestibulocerebellum (Gould and Butcher, 1987). Pseudo-cholinesterase was localized in adult Purkinje cells of the 23

Ch. I

J. Voogd, D. Jaarsma and E. Marani

lobules IX and X (Robertson et al., 1991). These cells are arranged in multiple, sagittal bands (Gorenstein et al., 1987). Robertson et al. (1991) were unable to confirm the transient staining with AChE in rat Purkinje cells. Somatostatin was located in rat Purkinje cells using polyclonal and monoclonal antibodies against conjugates of somatostatin (Johansson et al., 1984; Vincent et al., 1985; Villar et al., 1989). Reactive Purkinje cells were especially numerous in parts of the vermis and paraflocculus and flocculus during early postnatal stages, but mostly disappeared later on (Figs 19 and 20). In part of the vermis they were located in bands. In the adult rat Purkinje cells can be stained on the ventral aspect of the paraflocculus (Gonzalez et al., 1988) and in the vermis, after interventricular administration of colchicine (Villar et al., 1989). Somatostatin-like immunoreactivity was also observed in climbing fibers, that were correlated with the patches of immunoreactive Purkinje cells and, more diffusely distributed, in Golgi cells (Villar et al., 1989). The presence of somatostatin in adult rat Purkinje cells of the paraflocculus was confirmed with nonradioactive in situ hybridization for somatostatin mRNA (Kiyama and Emson, 1990). Specific binding of iodinated agonists of somatostatin was studied in rat, using ligands for short, 14 amino-acid ([125I]SS-14) and long forms ([125I]SS-28). Binding in the cerebellar cortex was found to be low, but strong binding of both ligands was observed over the cerebellar nuclei (Uhl et al., 1985). Binding to somatostatin receptors in the human cerebellar cortex was higher. Different distribution patterns were noted among the patients studied, with higher densities over the granular layer (Laquerriere et al., 1994). Leroux et al. (1985) and Gonzalez et al. (1988) failed to demonstrate specific binding over the cerebellar nuclei of the rat of a different SS-14 ligand, but confirmed binding of SS-28 (Leroux et :al., 1985). Binding of an octopeptide somatostatin analogue was reported to be almost absent in rat cerebellum (Reubi and Maurer, 1985) and low over the cerebellar cortex of the human cerebellum, with intermediate values in the molecular layer (Reubi et al., 1986). Tyrosine hydroxylase, the synthesizing enzyme of dopamine, is expressed by Purkinje cells of the ventral vermis (lobules I and X) and the hemisphere (ansiform lobule, paraflocculus) of rat cerebellum (Takada et al., 1993). Expression of tyrosine hydroxylase by Purkinje cells is increased in the mutant tottering and leaner mice (Austin et al., 1992). 3.1.4. The localization of the IP3 receptor and the intracellular calcium stores of Purkinje cells

The phosphoinositide system is a second messenger system coupled to metabotropic, G protein-linked receptors (see Ross et al. (1990), Mayer and Miller (1990), Ferris and Snyder (1992) and Berridge (1993), for reviews). Receptor-mediated hydrolysis of phosphatidylinositol (PIP2) is catalyzed by phospholipase C and leads to the formation of inositol-l,4,5-triphosphate (IP3) and diacylglycerol (DAG), two second messengers that function in a bifurcating signal pathway. Other inositol phosphates (inositol 1,3,4,5tetrakiphosphate, IP4; inositol 1,3,4,5,5-pentakiphosphate, IPs; and inositol hexakiphosphate, IP6) have been localized in rat cerebellum (Vallejo et al., 1987; Theibert et al., 1987, 1991). Phosphorylation of IP 3 by the enzyme IP 3 3-kinase leads to the formation of IP4. IP3, through activation of IP3 receptors, causes Ca 2+ mobilization from intracellular sources, whereas DAG, together with Ca 2+, activates the enzyme protein kinase C that phosphorylates regulatory proteins. The localization of phospholipase C, IP 3 receptors and protein kinase C has been extensively studied in Purkinje cells. 24

The cerebellum: chemoarchitecture and anatomy

Ch. I

B

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f

Fig. 19. Schematic illustration of the zonal distribution of somatostatin immunoreactive Purkinje cells at

different levels of the cerebellum of a 21 day old rat. Drawings have been made from frontal, cresyl-violet stained sections. Each dot represents 2-5 cells. Abbreviations: 5-9, cerebellar lobules V-IX; 4V, 4th ventricle; COP, copula pyramis; CR2, crus 2, ansiform lobule; FL, flocculus; PFL, paraflocculus; PM, primary fissure; SF, secondary fissure. Villar et al. (1989).

IP 3 3-kinase, the enzyme that produces IP 4 from IP3, was exclusively localized in Purkinje cells of the rat using immunohistochemistry (Mailleux et al., 1991 a, Mizuguchi et al., 1992) and in situ hybridization in rat and human cerebellum (Mailleux et al., 1991 b, 1992). Immunoreactivity was present in Purkinje cell dendrites more than in the perikarya. Intense immunolabelling of the dendritic spines was observed in the rat (Yamada et al., 1992; Go et al., 1993) (Fig. 21) but a specific role of IP 4 in Purkinje cell dendritic spines has not been disclosed. A similar localization in Purkinje cell dendritic spines was described for the mGluR1 subunit of the metabotropic glutamate receptor (Section 3.3.2., Fig. 52). Different isoenzymes of the phospholipase C (PLC) family, belonging to three major groups (fl, ~ and d), have been identified (Rhee et al., 1989; Rhee and Choi, 1992). PLC-fll, PLC-y and PLC-~ have been localized with in situ hybridization in the brain of the rat. Moderate activity was found for PLC-fll in the granular layer and strong activity in Purkinje cells and granule cells for PLCT'. The activity of PLC-d is low and may be localized in glial cells (Choi et al., 1989). PLC-A m R N A that was localized in rat Purkinje cells by Ross et al. (1989b), probably codes for a thiol-protein disulphide oxido-reductase and not for a PLC (Berridge, 1993). The IP 3 receptor has been found to be identical to the Purkinje cell-specific P400 25

Fig. 20. Examples of somatostatin-immunoreactive elements in the cerebellar cortex of the paraflocculus of adult colchicine treated rats. Patch of Purkinje cells and an immunostained Golgi cell (arrow head) are present in this section. Somatostatin-imrnunoreactive climbing fibers are observed. Calibration bar 50 prn. Villar et al. (1989).

5

The cerebellum." chemoarchitecture and anatomy

Ch. I

protein (Mignery et al., 1989) (Fig. 22). The P400 protein was originally isolated by Mallet et al. (1976) as a Purkinje cell-specific protein, that was reduced in homozygous Purkinje cell-deficient (pcd, Mullen et al., 1976) and 'staggerer' (Sidman et al. 1962) mice, but relatively enriched in the cerebella of 'reeler' and 'weaver' mutant mice, with a loss of granule cells (Mikoshiba et al., 1979). Immunocytochemical studies with a monoclonal antibody specific for P400 protein, indicated that the protein was localized in somata, dendrites and axons of Purkinje cells in rodents (Maeda et al., 1988; Nakanishi et al., 1991; Rodrigo et al., 1993). The development of Purkinje cells could be traced with P400-immunostaining of staged cerebella of mouse embryos (Maeda et al., 1989) (Fig. 24). At the ultrastructural level it was identified on the plasma-membranes and the endoplasmatic reticulum, including the subsurface cisterns (Maeda et al., 1989). Notably Purkinje cells of'staggerer' mice, that are defective in synaptic contacts of parallel fibers and lack dendritic spines, do not express P400-immunoreactivity, whereas P400-immunoreactivity was found at 'normal' levels in ectopic Purkinje cells of 'reeler' cerebellum (Mariani et al., 1977; Mikoshiba et al., 1980; Maeda et al., 1989) (Fig. 23), and in the few remaining Purkinje cells of 'pcd' mutant mice. Cloning of the P400 protein cDNA revealed that it was identical to the IP 3 receptor protein, as well as the Purkinje cell-specific PCPP-260 protein isolated by Walaas et al.

Fig. 21. IP3-3-kinase immunoreactivity in the rat cerebellum. Electron micrograph showing intense immunoreactivity in the dendritic spines of Purkinje cells. Bar 2 r Yamada et al. (1992).

27

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J. Voogd, D. Jaarsma and E. Marani

Fig. 22. Localization of inositol 1,4,5-triphosphate receptor with PCD6 antibody in frozen sections of rat cerebellum by immunofluorescence. Sagittal section of the cerebellar cortex. Small arrows in the granule cell layer (GL) point to segments of immunoreactive axons which represent recurrent collaterals of Purkinje cell axons. Mignery et al. (1989).

(1986) and the PDC6 protein of Nordquist et al. (1988). The localization of P400 (= IP3 receptor) mRNA in Purkinje cells was confirmed by in situ hybridization (Furuichi et al., 1989) (Fig. 25). The IP3 receptor was purified from rat cerebellum as a protein with a molecular weight of 260 kDa (Supattapone et al., 1988). The primary structure of the mouse IP3 receptor protein, and its partial homology to the skeletal muscle ryanodine receptor were elucidated by Mignery et al. (1990). The IP3 receptor is composed of four identical subunits of a molecular weight of 320 kDa, and forms a calcium-permeable channel (Maeda et al., 1991). Three additional cDNAs encoding for the IP3 receptor, 28

The cerebellum." chemoarchitecture and anatomy

Ch. I

named IP~R-II, III and IV, were identified by Sfidhof et al. (1991) and Ross et al. (1992), but were not found to be expressed at significant levels by Purkinje cells. The presence of the IP~ receptor in Purkinje cells was confirmed immunocytochemically. In immunocytochemical studies with gold-conjugates, that allow precise ultrastructural localization of the immunoreactivity, it was shown that gold particles were located on membranes of the endoplasmatic reticulum in somata, dendrites, dendritic spines and axons of the Purkinje cells (Mignery et al., 1989; Ross et al., 1989a; Sharp et al., 1993a,b) (Fig. 26). Immunolabelling predominated in the smooth-surfaced endoplasmatic reticulum, including the subsurface cisterns, but was also found on portions of the perinuclear and rough endoplasmatic reticulum, and on the cis-cisternae, but not the intermediate and trans-cisternae, of the Golgi apparatus. IP~-receptor immunoreactivity was also observed in a subpopulation of spherical or elongated, membrane-bound structures, named calciosomes (Volpe et al., 1989), that are present throughout the cytoplasm of the Purkinje cells (Volpe and Villa, 1991; Nori et al., 1993). Strong immunoreactivity for the IP~ receptor was found on stacks of flattened cisternae of the endoplasmatic reticulum (Otsu et al., 1990; Satoh et al., 1990; Takei et al., 1992, 1994). The labelling on the cisternal stacks was mostly located in the spaces between the cisternae and between the cisternae and the plasmalemma or mitochondria (Satoh et al., 1990; Takei et al., 1992, 1994). It should be noted that the amount of cisternal stacks in Purkinje cells may depend on the conditions of perfusion fixation. The presence of cisternal stacks in healthy Purkinje cells, therefore, has been disputed (Takei et al., 1994).

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Fig. 23. Section of reeler m u t a n t m o u s e cerebellum stained with m o n o c l o n a l a n t i b o d y 4C11 against the P400 protein. N o t e stained Purkinje cells in the cortex (CX) and in the central mass o f dislocated cells (DP). Bar = 200/~m. M a e d a et al. (1989).

29

Ch. I

J. Voogd, D. Jaarsma and E. Marani

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Fig. 24. Sagittal sections of mouse cerebella of various ages stained with a monoclonal antibody (4C 11) against the P400 protein.The samples were from (A) postnatal day O (PO), (B) P3, (C) P5, (D) P7, (E) P10, (F) P15, and (G) P20 cerebellum. A section from a P20 old mouse cerebral cortex did not react with same antibody (H). Magnification 100• Maeda et al. (1989).

30

The cerebellum." chemoarchitecture and anatomy

Ch. I

Nevertheless the formation of stacks of cisternae of the endoplasmatic reticulum could be induced by overexpression of IP3 receptors in fibroblasts, which indicates that cisternal stacks may exist as special organelles related to the IP3 receptor (Takei et al., 1994). The localization of the IP3 receptor has been compared to the localization of other luminal or membrane components of the endoplasmatic reticulum related to Ca 2+ homeostasis. The membrane pump CaZ+-ATPase, immunolabelled with antibodies against cardiac CaZ+-ATPase, was found to be located in regular cisternae of the endoplasmatic reticulum, the lateral tips of cisternae of the Golgi complex and in calciosomes of Purkinje cells (Kaprielian, 1989; Michelangeli et al., 1991; Villa et al., 1991; Takei et al., 1992) (Fig. 27). Distal axons of Purkinje cells, however, lacked CaZ+-ATPase immunoreactivity (Takei et al., 1992). Appreciable levels of calsequestrin, the main intraluminal calcium-binding protein of muscle, were present in Purkinje cells of the chicken. Calsequestrin-immunoreactivity was present over the lumen (Villa et al., 1991) and membranes (Takei et al., 1992) of stacked and isolated cisternae of the endoplasmatic reticulum and in a subpopulation of calciosomes (Volpe et al., 1988; Volpe and Villa, 1991). Mammalian Purkinje cells do not have calsequestrin but, instead, express calretic-

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Fig. 25. Localization of the P400-specific mRNA by in situ hybridization, a. Autoradiograph of a sagittal section of mouse cerebellum, b. Higher magnification of a. ML, molecular layer; PL, Purkinje cell layer; GL, granular layer. Furuichi et al. (1989).

31

Ch. I

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J. Voogd, D. Jaarsma and E. Marani

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Fig. 26. Electron-microscope immunocytochemical localization of InsP3 receptor in Purkinje cells of rat cerebellum using pre-embedding avidin-biotin labelling and InsP3 receptor antiserum. A, B. Nuclear membrane and some, but not all, portions of endoplasmic reticulum (ER) are labelled. C. Higher magnification of the dendritic pole of a labelled Purkinje cell. Note the unlabelled ER very close to labelled ER. D. Labelled portions of endoplasmic reticulum (L-ER) immediately subjacent to an unlabelled presynaptic terminal (U-T). Cell membrane indicated with triangles in (D) and (E). E. InsP3 receptor antiserum. Labelled portions of ER near plasma membrane, but not directly subjacent to presynaptic terminal. F. Preimmune serum. No specific label is present, even though the section is very close to the surface of the vibratome section. Abbreviations: L-ER, labelled endoplasmic reticulum; L-G, labelled Golgi apparatus; L-NM, labelled nuclear membrane; U-ER, unlabelled endoplasmic reticulum; U-G, unlabeled Golgi apparatus; U-M, unlabelled mitochondrion; U-NM, unlabelled nuclear membrane. Scale bars for all panels 1 r (Ross et al., 1989a).

ulin (Treves et al., 1990). Calreticulin-immunoreactvity was located in stacks of rough and smooth endoplasmatic reticulum in rat Purkinje cells (Nori et al., 1993). Calsequestrin and calreticulin are not exclusively present in Purkinje cells, but also in other cell types of the cerebellar cortex. 3.1.5. Protein kinase C in Purkinje cells

Protein kinase C (PKC) plays an important role in the control of several cellular processes, such as the short-term modulation of membrane excitability and transmitter release, positive or negative interaction with the conductance through various ion channels and the regulation of gene expression and cell proliferation (Shearman et al., 1989, 1991; Farago and Nishizuka, 1990; Nishizuka et al., 1991). PKC, that through phosphorylates multiple target protein including neurotransmitter receptors, and has been implicated in long-term depression (LTD) of glutamate sensitivity of Purkinje cells (Cr~pel and Krupa, 1988). Breakdown of PIP2 by phospholipase C (see Section 3.1.4) in Purkinje 32

The cerebellum." chemoarchitecture and anatomy

Ch. I

a

b

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Fig. 27. Immunofluorescence localization of the cerebellar Ca2+-ATPase in a transverse cryosection of adult chicken cerebellum. CaS/CI-IgG localizes the Ca2+-ATPase to the Purkinje cell bodies in the Purkinje layer (b), and the dendritic trees in the molecular layer (a). Very faint immunofluorescence was detected in the granule cell layer (c). Bar 50/~m. Kaprielian et al. (1989).

cells can activate PKC through the production of DAG and the mobilization of C a 2+ from the endoplasmatic reticulum. Alternative routes for the production of DAG and the mobilization of C a 2+ from extracellular sources are available (Nishizuka et al., 1991). Three isoenzymes of PKC have been distinguished on the basis of the analysis of the sequence homology of complementary DNA clones from different sources. The PKCtypes I, II and III of Huang et al. (1987a,b) are the products of the 7', fl and ~ genes respectively (Ono et al., 1987; Nishizuka, 1988). The PKC fl isoenzyme occurs in two forms, flI and flII, generated through alternative splicing (Ono et al., 1987; Nishizuka, 1988; Saito et al., 1989; Shimohama et al., 1990; Farago and Nishizuka, 1990). PKC ~, fl and 7' are calcium-dependent forms. In addition, calcium-independent isoenzymes of PKC have been identified: ~, e, e' and ( (Ono et al., 1988). Non-specific antibodies against PKC were found to strongly immunostain Purkinje cell perikarya, dendrites and axons (Mochly-Rosen et al., 1987; Kitano et al., 1987; Saito et al., 1988). Immunocytochemical studies with subtype specific antibodies and in situ hybridisation histochemistry have shown that several PKC subtypes are located in Purkinje cells (Figs 28 and 29, Table 1) (Brandt et al., 1987; Huang et al., 1987a,b, 1988, 1991; Ase et al., 1988; Hashimoto et al., 1988; Hidaka et al., 1988; Kose et al., 1988; Shimohama et al., 1990; Wetsel et al., 1992; Chen and Hillman, 1993a; Garcia et al., 1993; Merchenthaler et al., 1993). PKC~' immunoreactivity occurs at high levels in both the somatodendritic and axonal domains of Purkinje cells, and is absent from other cell types of the cerebellar cortex. Immunoreactivity for PKC ~ is also present in Purkinje 33

Ch. I

J. Voogd, D. Jaarsma and E. Marani

cells. PKC d-immunoreactive Purkinje cells are distributed in immunopositive and immunonegative columns (Fig. 133) (Chen and Hillman, 1993a). According to Wetsel et al. (1992) Purkinje cells were stained with antisera against PKC e, but Chen and Hillman's (1993a) found Purkinje cells to be unlabelled for PKC e. PKC/6 and e' were not located in Purkinje cells (Table 1). 3.1.6. cGMP; cGMP-dependent protein kinase and nitric oxide synthase in Purkinje cells Purkinje cells are the only cerebellar cell type containing cyclic guanosine 3',5'-monophosphate (cGMP)-dependent protein kinase (cGK) (Walter et al., 1981; Walter, 1984; Lohmann et al., 1981; De Camilli et al., 1984; Wassef and Sotelo, 1984). cGK-immunoreactivity is present throughout the entire Purkinje cell, including its dendrites and its axon (Fig. 30). During development Purkinje cells display a heterogeneity in their expression of immunoreactivity for cGK (Wassef and Sotelo, 1984, rat; Levitt et al., 1984, monkey) (see Section 6.2.). A 23 kD protein, which is likely to be a substrate of cGK was found to be concentrated in Purkinje cells (Walter, 1984; Nairn and Greengard, 1983). Immunoreactivity for guanylate cyclase, the synthesizing enzyme of cGMP, was

Fig. 28. Developmental expression of protein kinase C (PKC) isoenzymes in rat cerebellum. Immunofluorescent staining of cerebellar cortex by antibodies specific for PKC 1, corresponding to PKC~" (panels A, B and C), PKCfl (panels D, E and F) and PKC~ (panels G, H and I). Sagittal sections of cerebellum of 1-week-old (A, D and G), 2-week-old (B, E and H) and 3-week-old (C, F and I) rats were used. PKCz- antibody stained mainly the Purkinje cell bodies and dendrites throughout the development. PKCfl antibody stained the cerebellar granule cells in the external germinal layer (EGL) of the 1- and 2-week-old rats and mainly the granular layer of the 3-week-old rats. PKC~ antibody stained both granule cells and Purkinje cells throughout the development. Huang et al. (1991).

34

The cerebellum." chemoarchitecture and anatomy

Ch. I

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.

.

.

.

..

.

9

"

Fig. 29. Immunostaining for different isozymes of PKC in the rat cerebellum. PKC0~ (A) is present in Purkinje cells (P). The dendrites of these cells can be followed as far as the top of the molecular layer (m). The granular layer (g) and the white matter (w) are not stained. To avoid crowding, the abbreviations for layers of the cerebellum are indicated only in (A); however, the layer of Purkinje cells (P) is indicated in each figure for orientation. PKCfl (B) and PKCflI I (C) are present only in cells of the granular layer. PKCg is present in Purkinje cells and Bergmann glial cells in the molecular layer (D). Not only the perikarya but also the dendrites of Purkinje cells in the molecular layer and their axons in the granular layer are immunopositive. The antiserum for PKC~ stained Purkinje cells (E) and presently unidentified cells below the unstained Purkinje cells (F). The dorsally located folia contain mainly unstained Purkinje cells. Their axonal origin is surrounded by immunopositive cells. In the basal folia, the Purkinje cells are immunostained. PKCe is present in Purkinje cell (G), whereas PKCe' is present in cells in the molecular and granular layers and in the nerve fibers surrounding the unstained Purkinje cells (H). Antiserum against PKC~" stained only Purkinje cells in the cerebellum (I). Bar 100 •m. Wetsel et al. (1992).

35

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J. Voogd, D. Jaarsma and E. Marani

localized in Purkinje cells, but does also occur in other cell types of the cerebellar cortex (Zwiller et al., 1981; Ariano et al., 1982; Nakane et al., 1983; Poegge and Luppa, 1988). cGMP was, however, found to be absent from rat Purkinje cells, using antibodies against conjugates of cGMP in combination with sodium nitroprusside-stimulation of cGMP synthesis. Prominent cGMP-immunoreactivity within the molecular layer was detected in Bergmann glial cells (Fig. 56) (Berkelmans et al., 1989; De Vente et al., 1989, 1990). Soluble guanylate cyclase is activated by nitric oxide (NO) (see Section 3.4). NO has been implicated in the generation of long term depression (LTD) of parallel fibermediated EPSP's in Purkinje cells. LTD can be prevented by the application of haemoglobin that absorbs NO, or by the inhibition of NO synthesis (CrGpel and Jaillard, 1990; Shibuki and Okada, 1991; Ito, 1991). However, nitric oxide synthase, the synthesizing enzyme of NO, appears to be absent from the Purkinje cell (Section 3.4.).

3.1.7. Calcium-binding proteins in Purkinje cells Calbindin-D28K, parvalbumin and calmodulin are cytosolic, calcium-binding proteins of the EF-hand family (see Baimbridge et al., 1992 and Andressen et al., 1993 for reviews), that are present in high amounts in Purkinje cells. Calretinin is a calciumbinding protein closely related to calbindin-D28K, that is absent from the Purkinje cells, but present in other neurons and in afferent mossy and climbing fibers of the cerebellar cortex (Rogers, 1989; Arai et al., 1991; RGsibois and Rogers, 1992; Floris et al., 1994). One of the calcium-binding proteins, the 28 kDa vitamin-D-dependent calcium-binding protein (calbindin-D28K), occurs in most, if not all, Purkinje cells in rat and chicken cerebellum (Lawson, 1981; Roth et al., 1981; Jande et al., 1981a,b; Baimbridge and Miller, 1982; Legrand et al., 1983; Schneeberger et al., 1985; Kosaka et al., 1993; Amenta et al., 1994). Its presence in soma, dendrites and axon was demonstrated with polyclonal and monoclonal antibodies raised against calbindin-D28K (Fig. 31A). Its exclusive presence in the cerebellum in Purkinje cells was confirmed with in situ hybridization with cDNA probes in rat and mouse (Nordquist et al., 1988; Iacopino et al., 1990; Abe et al., 1992a; Kadowaki et al., 1993). According to Garcia-Seguera et al. (1984) only 74% of the rat Purkinje cells was immunoreactive for a polyclonal antibody raised against chick duodenal calbindin-28K. This antibody also stained Golgi cells in the granular layer in rat and human cerebellum (Fournet et al., 1986). Developmental gradients in the expression of immunoreactivity for calbindin-28K by Purkinje cells were studied by Legrand et al. (1983) and Wassef et al. (1985) (see Section 6.2.). TABLE

1. Immunoreactivities o f P K C in cerebellar neurons

Isoenzymes

P u r k i n j e cells

Basket &

G r a n u l e cells

Cerebellar nuclei

+

++

+

s t e l l a t e cells Alpha

++

Beta

-

+

++

+

Gamma

+++

-

-

-

Delta

+++

+++

-

-

Epsilon

-

+

++

++

Zeta

++

+

++

++

Chen and Hillman (1993a)

36

The cerebellum." chemoarchitecture and anatomy

Ch. I

P 21

Fig. 30. A. Frontal section through the cerebellum and attached brainstem of an adult rat. All the Purkinje cells are stained by cyclic 3',5'-guanosine monophosphate-dependent protein kinase (cGK) antiserum, including their dendrites in the molecular layer and their axon terminals in the deep nuclei and in the brainstem (arrow). Bar = 1 mm. B. Higher magnification of the neurons indicated by an arrow head in A. Like a few other isolated labelled cells found in variable locations, these cells are considered as ectopic Purkinje cells. Bar = 50/lm. C. cGK immunoreactive neuron in the cerebellum of 1 day-old rat. This ectopic Purkinje cell is located in the white matter and its appearance mimics that of 1-day-old Purkinje cells as visualized in Golgi impregnated material. Bar = 25 ~m. Wassef and Sotelo (1984).

Calmodulin-immunoreactivity was observed both in Purkinje cells and in cells of the cerebellar nuclei of the rat (Lin et al., 1980; Means and Dedman, 1980; Seto-Oshima et al., 1983, 1984). During postnatal development calmodulin-immunoreactivity was transiently present in the inner part of the external germinative layer and in fibers in the white matter of P3-P11 rat pups (Seto-Oshima et al., 1984). Polyclonal antibodies against parvalbumin stain all Purkinje cells and stellate and basket cells in the molecular layer of rat and avian cerebellum (Figs 31B and 32) (Celio and Heizmann, 1981; Heizmann, 1984; Braun et al., 1986; Endo et al., 1985; Schneeberger et al., 1985; Seto-Oshima et al., 1983; Rogers, 1989; Kosaka et al., 1993). The localization of parvalbumin in Purkinje, stellate and basket cells was confirmed in the rat with non-radioactive in situ hybridization (Kadowaki et al., 1993). Parvalbumin 37

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J. Voogd, D. Jaarsma and E. Marani

. .

Fig. 31. A. Calbindin-D28k immunoreactivity. B. Parvalbumin-immunoreactivity in rat cerebellar cortex. Purkinje cells react with both antibodies; arrows in B indicate parvalbumin- immunoreactive stellate and basket cells. Bar - 50/zm. Courtesy of Dr. M.P.A. Schalekamp.

supposedly occurs in GABAergic neurons (Celio and Heizmann, 1981) and/or neurons with characteristically high firing rates (Karmy et al., 1991). Karmy et al. (1991) studied the co-localization of parvalbumin and cytochrome oxidase, as an indicator of metabolic activity, in many regions of the brain. They found only weak immunoreactivity with antibodies against cytochrome oxidase in parvalbumin immunoreactive Purkinje cells of the rat. A developmentally regulated polypeptide (PEP-19), that is a presumptive neuronspecific calcium binding protein, was identified in adult and neonatal rat cerebellum and its amino acid sequence was determined (Ziai et al., 1986). PEP-19-like immunoreactivity is expressed by Purkinje cells and by the cartwheel cells of the dorsal cochlear nucleus of the mouse (Mugnaini et al., 1987). Berrebi et al. (1991) drew attention to the expression of PEP-19, CaBP and other Purkinje cell markers (cerebellin, L7: see below) by bipolar cells and other neurons of the retina.

3.1.8. Other specific biochemical markers for Purkinje cells Several polypeptides, that are present in all Purkinje cells, but not in other cells of the cerebellum, have been mentioned in the previous sections of this chapter. They include the IP3 receptor (identical to the P400 protein and to the PCPP-260 protein of Walaas et al., 1986) (see Section 3.1.4), IP3-3-kinase (Section 3.1.4), cGMP-dependent protein kinase (Section 3.1.5), PEP-19 and calbindin-D28K (Section 3.1.7). Two other 38

The cerebellum." chemoarchitecture and anatomy

Ch.I

proteins, cerebellin and L-7 that occur in all Purkinje cells, are dealt with in this section. Other proteins only occur in certain subpopulations of Purkinje cells. Zebrin I and II (Hawkes et al., 1985) are the prototypes of this group. The restriction of the Zebrins to a subpopulation of Purkinje cells is the more remarkable because they are originally present in all Purkinje cells of rat neonates (Leclerc et al., 1988). The developmental histories of cGMP-dependent protein kinase, calbindin-D28K and L-7 are quite different, in that these proteins make their first appearance in subpopulations of fetal Purkinje cells and only in later stages become expressed by all Purkinje cells of the cerebellum (Wassef and Sotelo, 1984; Smeyne et al., 1991) (Section 6.2.). Purkinje cell-specific markers include several glyco- and phosphoproteins, peptides, antigenic determinants that have not been identified or determinants that Purkinje cells share with other, non-cerebellar cell types. One of the first sera specific for rat Purkinje cells was obtained, using immunohistochemical screening, by Woodhams et al. (1979), but the antigen corresponding to this antibody has not been identified. Reeber et al. (1981) isolated a Purkinje cell specific 24 kDa glycoprotein from rat, that was present (Reeber et al., 1981) throughout the whole somatodendritic extent of the Purkinje cells, associated with the plasma membrane, as well as with the rough endoplasmatic reticulum and polysomes, the cytoplasmic side of the nuclear envelope and subsurface cisterns (Langley et al., 1982). Visinine, a soluble, 24 kDa protein, isolated from chicken retina, was found to be an exclusive marker for Purkinje cells in rat cerebellum (Yoshida et al., 1985). Specific staining of Purkinje cells was also found with monoclonal antibodies directed against human T cells (Garson et al., 1982), against certain cytoplasmic antigens in Purkinje cells (Weber and Schachner, 1982) and against antigenic determinants on trypanosomes (Wood et al., 1982). One of the antibodies (UCHT 1), isolated by Garson et al. (1982) is remarkable because its antigen is not present in Purkinje cells from 'staggerer' mutant mice (Caddy et al., 1982), a property the UCHT 1 antigen shares with the IP3 receptor protein (Section 3.1.4). One group of Purkinje cell-specific markers, the cerebellins, has been studied in more detail. A Purkinje cell-specific hexadecapeptide called 'cerebellin' and its metabolite, des-Serl-cerebellin were isolated and sequenced by Slemmon et al. (1984). Cerebellin immunoreactivity as studied with polyclonal antibodies in rat was found in soma and dendrites of nearly all Purkinje cells, but was absent beyond the initial axon segment (Slemmon et al., 1984). Cerebellin-immunoreactivity could also be demonstrated in cerebella of different species, including human and chick (Morgan et al., 1988), and in cartwheel cells and basal dendrites of pyramidal neurons of the dorsal cochlear nucleus (Fig. 33) (Mugnaini and Morgan, 1987). Cerebellin differs from most other markerproteins of Purkinje cells in being absent from other sites in the CNS, including the retina (Berrebi et al., 1991). Slemmon et al. (1988) and Morgan et al. (1988) concluded from an analysis of cerebellin immunoreactivity in Purkinje cells of different mutant mice with a varying loss of the granule cells, that the amount of cerebellin is correlated with the formation and the number of parallel fiber-Purkinje cell synapses. L-7 is a protein specific for Purkinje cells. Labelling with polyclonal antibodies against predicted L-7 sequences was present in somata, including the nucleus, in dendrites and dendritic spines, and in axon and axon terminals of Purkinje cells. All Purkinje cells, but no other types of cerebellar neurons appeared to be labelled (Berrebi and Mugnaini, 1992). The expression of the L-7 gene by all adult Purkinje cells of the rat cerebellum was reported by Nordquist et al. (1988, their PCD5 clone), Oberdick et al. (1990) Vandaele et al. (1991, their Purkinje cell protein-2) and Smeyne et al. (1991). According to Oberdick et al. (1990) and Berrebi et al. (1991) the L-7 gene is also expressed by retinal 39

Ch.I

J. Voogd, D. Jaarsma and E. Marani

Fig. 32. Parvalbumin immunoreactivity in the developing cerebellar cortex of the zebra finch. A. Incubation day D 16: Clusters of labelled Purkinje cells of varying staining intensity. Stained Purkinje cells axons are seen in the internal granular layer (IGL). Note the areas containing unstained or only slightly stained cells and the dot-like staining pattern in the external granular layer (EGL). B. Adult: The dendrites of the Purkinje cells have reached the cerebellar surface and are now fully branched. Between them many immuno-stained basket and stellate cells are visible. Parvalbumin immunoreactivity in Purkinje cell axons is no longer visible except for a few fragments lying in the internal granular layer (IGL). The layer of Purkinje cells is still interrupted by parvalbumin immunonegative areas. Calibration bar in A - 50/lm, in B = 100/~m. Braun et al. (1986).

bipolar cells. The initial expression of the L-7 gene by zonally distributed Purkinje cells during prenatal and early postnatal development was studied by Vandaele et al. (1991), Smeyne et al. (1991) and Oberdick et al. (1993) (see also Section 6.2.). 40

The cerebellum." chemoarchitecture and anatomy

Ch. I

Several other markers are only present in zonally distributed subpopulations of Purkinje cells (see also Section 6.1.3.). The monoclonal antibody B1 of Ingram et al. (1985) was raised against rat embryonic forebrain membranes. Purkinje cells in broad parasagittal bands, alternating with B 1-negative zones, were immunoreactive in the cerebellum of Macaca fascicularis. Other neurons in the molecular layer and cells of the cerebellar nuclei were also stained by this antibody. A similar pattern of B l-immunoreactivity was present in the cerebellum of the rat. The monoclonal antibody mabQ 113 was developed, specified and used in anatomical studies by Hawkes et al. (1985), Hawkes and Leclerc (1986, 1987), Hawkes and Gravel (1991), Hawkes (1992) and Leclerc et al. (1992). It is directed against a 120 Kda protein (Zebrin I); the function of this protein is still unknown. A specific subpopulation of Purkinje cells displays immunoreactivity for Zebrin I in their dendrites, soma, axon and axon terminals. Zebrin I-positive and negative Purkinje cells are distributed in parasagittal bands (Fig. 34) (see also Section 6.1.3.). Ultrastructurally Zebrin I-immunoreactivity in rat Purkinje cells is localized in the cytosol. It is absent from membrane-bound organelles such as the mitochondria and the synaptic vesicles. In large dendrites reaction product is associated with microtubuli, in spines it is located at the postsynaptic densities. An antibody raised against the cerebellum of the weakly electric fish Apteronotus (anti-Zebrin II: Brochu et al., 1990) recognizes the same Purkinje cells as anti-Zebrin I in the cerebellum of the rat, and is effective in staining these neurons in a large number of other species such as the opossum (Fig. 137). The epitope of the Zebrin II antibody is associated with a 36 kDa polypeptide identified as the glycolytic enzyme aldolase C. In situ hybridization of Zebrin II mRNA showed a strong signal in mouse Purkinje cells with normal regional heterogeneity (Hawkes, 1992; Ahn et al., 1994). Rat Purkinje cells containing low affinity nerve growth factor receptor protein (Sotelo and Wassef, 1991; Dusart et al., 1994) (see Section 3.1.10 and Fig. 38C,D), PKC delta (see Section 3.1.10 and Fig. 133), or the monoclonal antibody B30 of Stainier and Gilbert (1989), that recognizes two minor gangliosides, show the same distribution as Zebrin-stained Purkinje cells. Although the distribution of the enzyme 5'-nucleotidase in the molecular layer of rat and mouse cerebellum (Scott, 1963; Marani, 1982a,b) is identical to that of the Zebrins (Eisenman and Hawkes, 1993) (Fig. 135), it may be located in Bergmann glial and not in Purkinje cells (see Section 3.5.). Several proteins are distributed in more or less complementary patterns, either in Zebrin-negative Purkinje cells (Ppath, HNK, cytochrome oxidase) or in Bergmann glia (3a-fucosyl-N-acetyl lactosamine [FAL], glycogen phosphorylase). The antibody P-path is directed against acetylated gangliosides (Edwards et al., 1989, 1994; Leclerc et al., 1992) and reacts with Zebrin-negative Purkinje cells in mouse cerebellum (Fig. 134). The localization of cytochrome oxidase was described by Hess and Voogd (1986), Leclerc et al. (1990) and Harley and Biejalew (1992) in the cerebellum of macaques, the squirrel monkey and the rat. The localization of HNK was studied by Eisenman and Hawkes (1993) in the mouse. The FAL-epitope (Fig. 94; Bartsch and Mai, 1991) and the enzyme glycan phosphorylase (Marani and Boekee, 1973; Harley and Bielajew, 1992) have been located in subsets of mouse Bergmann glial cells, that are distributed in a complementary manner with respect to the Zebrin pattern. Gangliosides are glycolipids, concentrated in the outer layer of neural plasma membranes. Biochemical analysis showed a correlation between the selective degeneration of Purkinje cells in pcd and nervous mutant mice with the loss of the ganglioside GT~A. GT~A was more concentrated and the ganglioside GD~A was diminished in weaver mutant mice with a selective loss of the granule cells (Seyfried et al., 1983, 1987; Marani 41

Ch. I

J. Voogd, D. Jaarsma and E. Marani

Fig. 33. Light photomicrographs showing cerebellin immunoreactivity in rat cerebellum (A-C) and the dorsal cochlear nucleus (D-F) in parasagittal sections. A. Cerebellar hemisphere with part of the underlying dorsal cochlear nucleus (DCoN, arrowhead). CN, cerebellar nuclei. B. Immunostaining in DCoN. The cell bodies of cartwheel neurons in the superficial layers (layers 1 and 2) of the DCoN and the plexus in the deeper region (layer 3) predominate. The plexus is most dense in the upper portion of the deep region, which may correspond to layer 3 of the feline nuclei, a zone that contains the basal dendritic arbors of the bipolar pyramid neurons, one of which is indicated by an arrow. C. Immunoreaction product is present in Purkinje cell body and main dendrites. D. In the axon, immunostaining is restricted to the initial axon segment (arrowhead). E. Three subependymal displaced Purkinje cells in DCoN. Smaller cell bodies of several cartwheel neurons (arrowheads) are also shown. F. Portion of the ventral cochlear nucleus in which immunostaining is restricted to rare cartwheel cell bodies (arrowheads) displaced in the superficial granular layer. Bars in A and B = 0.5/~m, in C-F = 50 r Mugnaini et al. (1987).

a n d M a i , 1992). A n o t h e r g a n g l i o s i d e , GD3, was localized in i m m a t u r e P u r k i n j e cells o f the rat, u s i n g a m o n o c l o n a l a n t i b o d y (Fig. 35). I m m u n o r e a c t i v i t y d i s a p p e a r e d f r o m the cell b o d y in the adult, b u t r e m a i n e d p r e s e n t in the m o l e c u l a r layer ( R e y n o l d s a n d Wilkin,

42

The cerebellum." chemoarchitecture and anatomy

Ch. I

.,.....

.".: ,, ~

, ~ ;~,

'

; .

,

. -i:

;~:57

'";'

:'~'~":

'2., ~

Fig. 34. 50/lm horizontal sections through the cerebellar cortex of the rat at postnatal day 25 to show the distribution of mabQ113 (Zebrin I) immunoreactivity. A. The peroxidase reaction product is confined exclusively to a subset of Purkinje cells that are distributed symmetrically into parasagittal compartments in both the vermis and hemispheres. Labelling of the bands of Zebrin I-immunoreactive Purkinje cells P l+ to P7+ according to Hawkes and Leclerc (1987). Scale bar = 500/~m. B. A higher-power view of P5 + and P6 + of the posterior lobe hemisphere, in the lobules bordering the intercrural fissure. Immunoreactivity is seen to extend throughout the Purkinje cell, and no other cell types in the cerebellum are stained. Scale bar = 200/zm. C. In addition to the regular band display, additional narrow 'satellite' bands are also common. The arrowheads indicate two such satellites in the posterior lobe vermis. Scale bar = 100/lm. Leclerc et al. (1988).

1988). Levine et al. (1986), who used another monoclonal antibody against GD3, found immunoreactivity of reactive astrocytes in mouse mutants, but failed to observe a reaction within the Purkinje cells. These different results probably are due to differences in fixation (Reynolds and Wilkin, 1988). 3.1.9. Cytoskeleton and metabolism of Purkinje cells The DNA content of mature Purkinje cells is high. Feulgen-DNA or propidiumiodideDNA reveal hyperdiploid values (Bernocchi, 1986; Bernocchi et al., 1986). Purkinje cells stand out by their high content of enzymes, mostly dehydrogenases (Adams, 1965). Their content of the glycolytic enzyme enolase is low (Pelc et al., 1986; Vinores et al., 43

Ch. I

J. Voogd, D. Jaarsma and E. Marani

1984). However, Purkinje cells of the human cerebellum stand out from other nerve cells by their high content of aldolase-C (Royds et al., 1987). Purkinje cells do not react with antibodies against the phosphorylated forms of the 70, 150 and 200 kDa neurofilament proteins (Pelc et al., 1986; Matus et al., 1979; Marc et al., 1986; Langley et al., 1988). The phosphorylated form of the 200 kDa protein is present in axons in the granular layer, that were identified as Purkinje cell axons by Marc et al. (1986) and as mossy fibers by Langley et al. (1988), both in the rat (Fig. 36). The non-phosphorylated form of the neurofilament proteins was found to be present in the entire Purkinje cell with the exception of distal dendrites. According to Marc et al. (1986) the protein is present as filamentous aggregates. Langley et al. (1988) stated that a monoclonal antibody against the non-phosphorylated form of the 200 kDa protein is present in soma and dendrites as patches of diffuse immunoreactivity without a filamentous substructure. In Friedreich's ataxia neurofilament, mainly the phosphorylated form, is expressed by human Purkinje cells within their soma and dendrites (Marani, unpublished results) (Fig. 37). The process of endocytosis in Purkinje cell has been studied in relation to synaptogenesis of the Purkinje cell dendrites. Glycoproteins located on the parallel fiber are also pinocytosed into the Purkinje cell. Lysosomal action degradates these glycoproteins. In this process alpha-D-massosidase plays an important role, which is selectively present in the Purkinje cell dendrites (Dontenwill et al., 1983). Other glycoproteins, like K+Na+ATP-ase are not taken up, indicating a receptor-mediated recognition of some glycans of the glycoproteins. The specificity of the pinocytosis for certain molecules suggests that this recognition is the preliminary event in the establishment of Purkinje cell synapses.

3.1.10. Nerve growth factor and nerve growth factor-receptor protein in Purkinje cells Nerve growth factor-like immunoreactivity was present in Purkinje cell somata and dendrites, with dense labelling in the paraflocculus, and in neurons of the cerebellar nuclei and the lateral vestibular nucleus of rat cerebellum (Nishio et al., 1994). All but a few of the Purkinje cells of the adult rat cerebellum stain with an antiserum against basic fibroblast growth factor. Staining was observed in all cellular compartments (Matsuda et al., 1992). P75 nerve growth factor-receptor protein (NGF-R) is present in developing and adult Purkinje cells. Yan and Johnson (1988) and Cohen-Cory et al. (1989) described and reviewed the development of NGF-R in rat cerebellum. Low affinity NGF-R immunoreactivity has been demonstrated with species-specific monoclonal antibodies in Purkinje cells of adult rats (Pioro and Cuello, 1988, 1990; Pioro et al., 1991; Fusco et al., 1991; Dusart et al., 1994), monkey and human brain (Mufson et al., 1991). Immunoreactivity was present in the somata, dendrites and the proximal axon of the Purkinje cells. Additional immunoreactivity in granule cells was reported by Vega et al. (1994), using Bouin's fixative. NGF-R mRNA is expressed during early development in neurons of the rat external granular layer and in Purkinje cells. It peaks at postnatal day 10 and declines afterwards (Cohen-Cory et al., 1989; Lu et al., 1989) but also can be demonstrated in Purkinje cell somata in adult rodents (Fig. 38) (Koh et al., 1989) and primates (Mufson et al., 1991). NGF-R immunoreactivity was found to be highest in the flocculonodular lobe (Pioro and Cuello, 1988, 1990; Fusco et al., 1991). A distribution with strong expression in the flocculonodular lobe, the ventral parts of the anterior lobe and the lobules VII, VIII and 44

The cerebellum." chemoarchitecture and anatomy

Ch.I

Fig. 35. Double-immunofluorescent staining of 20-day rat cerebellar sections with antibodies to GD 3 ganglioside and glial acidic fibrillary protein (GFAP). Purkinje cell dendrites are intensely GD3-immunoreactive (A) but do not extend to the pial surface, unlike the Bergmann glial fibers (B), which project brightly GFAPimmunoreactive end-feet onto the pial membrane. Scale bar is 35 ~tm. Reynolds and Wilkin (1988).

45

Ch. I

J. Voogd, D. Jaarsma and E. Marani

..

9i:!?~iii~ &

1

F

.

-

.g:

!i . . . .

Immunocytochemical staining patterns of two monoclonal anti-bodies directed against nonphosphorylated and phosphorylated neurofilaments were studied in the cerebellum of developing normal rats. A. Non-phosphorylated neurofilaments on postnatal day 11. B. Day 21. Basket cell axons form a characteristic brush-like plexus around the initial segment of the Purkinje cell axon. C. Phosphorylated neurofilaments on postnatal day 13. D. Postnatal day 21. Stained filaments are restricted to Purkinje cell and basket cell axons and are absent from the Purkinje cell cytoplasm. Calibration bars in A and C 30/lm, in B and D 10 ~tm. Marc et al. (1986).

Fig. 36.

46

The cerebellum." chemoarchitecture and anatomy

Ch. I

IX of the caudal vermis and low activity in the hemisphere, was described by Mufson et al. (1991) for primates and man. The administration of colchicine results in the expression of N G F - R in most cerebellar Purkinje cells (Pioro and Cuello, 1988, 1990; Pioro et al., 1991). Koh et al. (1989) and Fusco et al. (1991) found N G F - R mRNA expression and NGF-R immunoreactivity in adult rat~ to be present in alternating Purkinje cell zones of strong and weak activity (Fig. 38C,D). This zonal pattern was also observed by Pioro and Cuello (1990). Its correspondence to the pattern of mabQ113 (Zebrin) immunoreactive zones (Hawkes and Leclerc, 1987) was noticed by Sotelo and Wassef (1991) and verified by Dusart et al. (1994) in adult rats. Lesions of the white matter, or knife cuts isolating the dorsal portion of the vermis of the rat cerebellum induces NGF-R immunoreactivity in previously unstained Purkinje cells (MartinezMurillo et al., 1993; Dusart et al., 1994).

3.1.11. Immunoreactivity of Purkinje cells in paraneoplastic diseases Specific forms of immunoreactivity of Purkinje cells have been discovered in human paraneoplastic conditions. Subacute cortical cerebellar degeneration in man may be associated with several types of carcinoma (see Vecht et al., 1991 for review). It has been most frequently observed in association with ovarian or endometrial carcinoma, but it also occurs as a rare sequal of small-celled bronchial carcinoma. It is generally characterized by a diffuse or patchy loss of Purkinje cells; granule cells also can be affected (Brain et al., 1951; McDonald, 1961; Brain and Wilkinson, 1965; Schmid and Riede, 1974; Steven et al., 1982). Strong labelling of Purkinje cells and weak staining of the granular layer was observed in sections of human cerebellum with a serum of patient with cerebellar degeneration with Hodgkin's disease using the indirect fluorescent staining procedure (Trotter et al., 1976). Sera of patients with carcinoma of the ovary were found to react with human Purkinje cells and neurons of the cerebellar nuclei using the same method (Greenlee and Sun, 1985). Jaeckle et al. (1985) distinguished a granular cytoplasmic and a diffuse form

9

.

.

.

. .

..

Fig. 37. Expression of phosphorylated neurofilament localization in normal human cerebellar cortex (A) and in Friedreich's disease (B). Note the strong positivity of the white matter and the molecular layers in a case of Friedreich's ataxia. No expression was found in the normal folium that was Nissl counterstained to demonstrate the granular and Purkinje cell layer. M = molecular layer, P = Purkinje cell layer, G = granular layer, F = fiber layer. Marani, unpublished.

47

Ch. I

J. Voogd, D. Jaarsma and E. Marani

Fig. 38. A. Nerve growth factor-R (NGF-R) transcripts are localized within Purkinje cells in the paraflocculus of rat cerebellum. B. NGF-R immunocytochemistry shows the perikarya of the Purkinje cells as well as the dense staining of the molecular layer, where the dendritic trees of the Purkinje cells arborize. Arrows in C and D point to parasagittal zones of intense labelling interdigitated with weaker labelling. Bar = 90 r Koh et al. (1989).

of immunoreactivity of human Purkinje cells with sera from patients with cerebellar degeneration suffering from ovarian or breast cancer. The diffuse form of Purkinje cell staining also was observed at higher concentrations with some sera of normal controls. Moreover the diffuse staining is not restricted to Purkinje cells, but also involves stellate, basket and some granule cells (Andersson et al., 1988). Cunningham et al. (1986) further analysed the sera causing granular deposits in the Purkinje cell cytoplasm with immunoblotting of extracts of human Purkinje cells. This so-called anti-Yo serum recognizes a 62 kDa and a 34 kDa protein. Antibodies raised against both proteins react with Purkinje cells in tissue sections (Fig. 39). The strongest reaction was observed for the antibody against the 62 kDa protein. The specificity of this reaction and the presence of anti-Yo immunoreactivity in tumor tissue was demonstrated by Furneaux et al. (1990). The 34 kDa antigen was found to correspond to the c D N A sequence of a clone recognized from a cerebellar expression library by a serum from a patient with paraneoplastic cerebellar degeneration (Dropcho et al., 1987; Furneaux et al. 1989). Other forms of immunoreactivity, with different cerebellar epitopes and a different localization of the immunoreactivity have been described (Tanaka et al., 1986; Smith et al., 1988; Rodriguez et al., 1988; Tsukamoto et al., 1989; Szabo et al., 1991). Differences in the localization of the immunoreactivity and in the characterization of the epitopes may be due to the use of rat cerebellum instead of human cerebellum in testing the sera by Tanaka et al. (1986), Smith et al., (1988) and Tsukamoto et al. (1989). Szabo 48

The cerebellum." chemoarchitecture and anatomy

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et al. (1991) isolated the (NuD) neuronal antigen recognized by sera from patients with paraneoplastic encephalomyelitis associated with small-celled bronchus carcinoma. This serum, also designated as anti-Hu, reacts with nuclei of neurons in the CNS, including the cerebellum (Andersson et al., 1988). 3.2. EXCITATORY PATHWAYS The cerebellar cortex is innervated by two types of excitatory afferents, the mossy and climbing fibers, and an intrinsic excitatory fiber system, the parallel fibers. An additional excitatory intrinsic pathway may be formed by unipolar brush cells, that give rise to mossy fiber-like fibers. The excitatory amino acid glutamate is the most likely neurotransmitter candidate for these pathways. An inherent problem in the localization of glutamate as a neurotransmitter is that there is no unequivocal marker for glutamatergic neurons and fibers since glutamate also participates in several metabolic pathways of nerve cells (Van den Berg and Garfinkel, 1971; Fonnum, 1984; Erecinska and Silver, 1990). The identification of glutamatergic pathways, therefore, is based upon a combination of anatomical, biochemical and physiological techniques (Fonnum, 1984). Immunocytochemistry with antibodies against glutamate (Storm-Mathisen et al., 1983) and physiological studies have proven to be particularly fruitful in the identification of glutamate as the neurotransmitter of the cerebellar excitatory pathways. These methods, however, do not totally exclude the possibility that other excitatory amino acids, such as aspartate or homocysteate, also participate as excitatory neurotransmitters. This holds in particular for the climbing fibers that have been frequently proposed to use aspartate as their primary neurotransmitter (see below). A major problem with 'nonglutamate' excitatory neurotransmitter candidates is that, as yet, no vesicular uptake

Fig. 39. Immunofluorescence of rat Purkinje cells with anti-Yo serum of a patient suffering from a cerebellar syndrome with ovarian carcinoma. Courtesy Dr. Ch. J. Vecht and Dr. J.W.B. Moll, Department of Neurology, Erasmus University Medical Center, Rotterdam.

49

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system has been isolated for these compounds. Instead, glutamate has been shown to accumulate in synaptic vesicles by a proton-driven vesicle transporter. This vesicle transporter is highly specific for glutamate, and in contrast to the cytoplasma membrane 50

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Fig. 40. Immunostaining in rat cerebellar cortex produced by anti-glutamate(Glu) mAb 2D7 (A,C,C',E and F) or by 'anti-GABA' mAb 3A12 (B and D). A, B (overview) and C, D (details) are from a pair of consecutive semithin sections. C and C' are enlargements of areas indicated in A and C, respectively. A and B. Note drastic difference in labelling patterns obtained with the two antibodies, gr, granule cell layer; P, Purkinje cell layer; mol, molecular layer. C. Frame indicating part of area shown enlarged in C'. C and D. Complementary labelling in stellate cells (stars), Golgi cell (arrows), pinceau formed by basket cell terminals (double arrow heads). Mossy fiber terminal-like structures in C (arrow head) fit into glomerular arrangements outlined by dots in D (arrow head). C'. Densely packed puncta probably represent parallel fiber terminals in molecular layer. E. Numerous strongly stained patches (arrow heads) and some fibers (arrow) are reminiscent of mossy fiber terminals. Granule cells with unlabelled nuclei appear less immunoreactive than those in C. F. Large mossy fiber terminal with several synaptic contacts (arrows) shows higher surface density of gold granules (EM immunogold procedure) than another terminal nearby (stars). Bars 100 ~tm in B, D and E, 1 r in F. Liu et al. (1989). (

transporter, does not transport aspartate (reviewed by Nicholls and Atwell, 1990; Jahr and Lester, 1992). 3.2.1. Mossy fibers Glutamate-like immunoreactivity in mossy fibers

Although subpopulations of mossy fibers may be peptidergic or cholinergic (see Sections 3.10. and 6.4.5.), it is now generally accepted that most if not all of the mossy fibers use L-glutamate as their principal neurotransmitter. The glutamatergic nature of mossy fibers has been evidenced with immunocytochemistry with antibodies against glutamateglutaraldehyde (Storm-Mathisen et al., 1983) or carbodiimide-glutamate conjugates (Madl et al., 1986). The rationale of this method is that glutamate, although ubiquiteously present throughout the neuronal cytoplasm at relatively high concentrations (~ 10 mM; Van den Berg and Garfinkel, 1971; Nichols and Attwell, 1990), is particularly enriched in glutamatergic nerve terminals, because of the presence of synaptic vesicles that concentrate glutamate to at least 60 mM. When electron microscopic post-embedding immunogold protocols are employed, quantitative and statistical analysis of the distribution of immunolabelling can be performed (e.g. see Ottersen, 1989). Glutamate immunoreactivity is widely distributed throughout the granular layer, but is enriched over mossy fiber rosettes in rat (Figs 40 and 41) (Ottersen and Storm-Mathisen, 1984a,b, 1987; Ottersen et al., 1987, 1990; Liu et al., 1989; Ji et al., 1991), cat (Somogyi et al., 1986) and monkey (Zhang et al., 1990). Mossy fiber rosettes contained significant higher levels of immunoreactivity than Golgi cell terminals and granule cell dendrites. Enriched glutamate-like immunoreactivity was also demonstrated in anterogradely horseradish peroxidase-wheat germ agglutinin (WGA-HRP) labelled spinocerebellar mossy fiber terminals. Notably, the density of glutamate-like immunoreactivity showed a strong positive correlation with the density of synaptic vesicles in these mossy fiber terminals (Ji et al., 1991). The anterogradely labelled mossy fiber terminals had a similar density of glutamate-like immunoreactivity as other mossy fiber rosettes. Mossy fiber terminals were not enriched in aspartate- or GABA-like immunoreactivities (Ji et al., 1991; Zhang et al., 1990). Data from physiological studies including recent patch-clamp studies are in line with the assumption that glutamate is the neurotransmitter of mossy fibers (Garthwaite and Brodbelt, 1989, 1990; Silver et al., 1992; D'Angelo et al., 1993; Rossi et al., 1995). 51

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Glutamine and glutaminase

Ottersen et al. (1992) quantified the compartmentalization of glutamate and glutamine in the cerebellar cortex of the rat, using post-embedding immunogold immunocytochemistry. They found the highest ratios of glutamate/glutamine in parallel fibers, high ratios in mossy and climbing fibers, low ratios in Purkinje and granule cells and in basket cell and Golgi cell terminals and the lowest ratios in Bergmann glia and astrocytes. This distribution is in accordance with uptake of glutamate from the extracellular space by glial cells, and its conversion into glutamine by the enzyme glutamine synthase, that is exclusively present in glia (Van den Berg and Garfinkel, 1971; Norenberg and MartinezHernandez, 1979; Fonnum, 1984; Erecinska and Silver, 1990). The glutaminase-glutamine loop is closed by diffusion of glutamine into neurons, that contain glutaminase, the enzyme that catalyzes the hydrolytic cleavage of glutamine to form glutamate. Wenthold et al. (1986) and Kaneko et al. (1987, 1989) used antibodies against glutaminase as an alternative approach to determine the cellular localization of glutamate. In the granular layer glutaminase-like immunoreactivity was present in granule cell somata (Wenthold et al., 1986) and in in small clusters, that probably represent mossy fiber rosettes (Fig. 42e) (Wenthold et al., 1986; Kaneko, 1987, 1989). Intense glutaminase-like immunoreactivity was also detected in several precerebellar nuclei, that give rise to mossy fibers, such as the pontine nuclei, the reticular nucleus of the pons, the lateral reticular nucleus, the vestibular nuclei and the external cuneate nucleus (Fig. 42a-d). Neurons in some of these nuclei have also been shown to react with antibodies against conjugates of glutamate (Beitz et al., 1986; Clements et al., 1986; Raymond et al., 1984). Glutamate transporters

The major mechanism by which synaptically released glutamate is inactivated is by highaffinity, sodium-dependent transport (Fonnum, 1984; Nicholls and Attwell, 1990). The sodium-dependent glutamate transporters are present in both neurons and astroglial cells, and have been assumed to be enriched on nerve terminals of glutamatergic axons. [3H]D-aspartate, a metabolically inert substrate of the glutamate transporter with very low affinity for glutamate receptors, has been widely used to locate glutamate or aspartate using fiber systems in the brain (Fonnum, 1984). Autoradiographic studies on cryostate sections indicate that [3H]D-aspartate binding sites are particularly enriched in the molecular layer, but are also present in the granular layer (Greenamyre et al., 1990; Anderson et al., 1990). Studies in cerebellar slices, however, show that [3H]Daspartate is not taken up by mossy fiber terminals (Garthwaite and Garthwaite, 1988). Accordingly, [3H]D-aspartate is not retrogradely transported by mossy fibers, allthough it is efficiently transported by climbing fibers (Wiklund et al., 1984). Three high-affinity sodium-dependent glutamate transporters have been cloned in rat: GLT-1 (Pines et al., 1992; Tanaka, 1993), EAAC1 (Kanai and Hediger, 1992), and GLAST (Storck et al., 1992). Recently, also four subtypes of human glutamate transporters, EAAT1-EAAT4, have been cloned with similar properties as their rat counterparts (Arriza et al., 1994; Fairman et al., 1994). In situ hybridisation and immunocytochemistry showed a differential distribution of the three transporters throughout the cerebellum. GLT1 is concentrated in the Bergmann glial fibers, but also occurs in the glial processes of the granular layer and in the cerebellar nuclei (Danbolt et al., 1992; Rothstein et al., 1994). High levels of GLAST are present in Bergmann glial fibers, but it is essentially absent from the granule cell layer. In the cerebellar nuclei it is mostly 52

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Fig. 41. Electron micrographs of serial sections through a glomerulus in the granular layer of the cat cerebellar cortex. The section shown in (A) was reacted with antiserum to glutamate (GLU), the section in (B) with antiserum to GABA. The electron-dense gold particles show immunoreactive sites. For GLU the highest density of gold appears to be over the mossy fiber terminal (mt) and the lowest over glial processes and Golgi cell terminals (1-3). This was confirmed by statistical comparison of the populations. The same Golgi cell terminals are strongly reacting for GABA, while other processes have only a low surface density of gold. Scale (A and B) 0.5/Ira. Somogyi et al. (1986).

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I

Fig. 42. Phosphate-activated glutaminase-like immunoreactivity (PAG-LI) in the precerebellar nuclei and the cerebellar cortex of the rat. Intensely labelled neuronal somata are seen in the pontine tegmental reticular nucleus of Bechterew (a), pontine nuclei (b), external cuneate nucleus (c), and lateral reticular nucleus of the medulla oblongata (d). Small clusters of grains, possible axon terminals, with PAG-LI are seen in the granular layer of the cerebellar cortex (e). Fine grains with PAG-LI are densely distributed, but no cell bodies are seen in the inferior olivary nucleus (f). CM, cerebellar medulla; G, granular layer; M, molecular layer; ML, medial lemniscus; P, pontine longitudinal fibers; Py, pyramidal tract; R, raphe. Scale bar 200 pm in a-d, f, 50 pm in e. Kaneko et al. (1987).

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associated with neurons (Rothstein et al., 1994). EAAC1 has been exclusively localized in neurons, with high densities in Purkinje cells and granule cells. Interestingly, immunocytochemical data show that EAAC1 is enriched in axon terminals of Purkinje cells, indicating that EAAC1 is not selective for glutamatergic nerve terminals. In accordance with the biochemical data, there was no immunocytochemical evidence for the presence of EAAC 1 or one of the other glutamate transporter proteins in mossy fiber terminals. Taken together the above data indicate that mossy fiber terminals are not provided with high-affinity glutamate transporters. Consequently, glutamate released by mossy fibers is likely to be predominantly cleared through glial cells (Wilkin et al., 1982; Garthwaite and Garthwaite, 1988). However, since glial processes do not enter the glomeruli (e.g. see Palay and Chan-Palay, 1974), an exclusive glial uptake implies that 'mossy fiber glutamate' molecules have to travel throughout extracellular space of the glomeruli before being inactivated. The clearance of 'mossy fiber glutamate' may be particularly slow at the giant mossy fiber-unipolar brush cell synapses, that may extend over 12-40 ,um2 with multiple clusters of presynaptic vesicles apposed to continuous regions of postsynaptic densities (Mugnaini and Floris, 1994). In fact, unusually long excitatory postsynaptic responses have been observed in unipolar brush cells following mossy fiber stimulation, consistent with a slow clearance of synaptically released glutamate (Rossi et al., 1995). 3.2.2. Climbing fibers Aspartate and glutamate

Several observations have led to the assumption that L-aspartate is the principal neurotransmitter of climbing fibers. (1) Destruction of the inferior olive in the rat with 3-acetylpyridine resulted in a small decrease in cerebellar aspartate concentration in total tissue homogenate (Nadi et al., 1977) and synaptosomal fractions (Rea et al., 1980). However, these observations were not confirmed by Perry et al. (1976). (2) It was demonstrated that after 3-acetylpyridine treatment Ca2+-dependent and K+-induced release of aspartate was significantly decreased (Toggenburger et al., 1983). Glutamate release was more dramatically decreased (e.g. see Cu6nod et al., 1989). (3) It was observed that climbing fibers but not mossy fibers in rat (Wiklund et al., 1984) and monkey (Matute et al., 1987) retrogradely transported [3H]D-aspartate. These experiments, however, only showed that climbing fibers are provided with high-affinity sodium-dependent glutamate transporter protein, and did not give information about the kind of transmitter used by the climbing fibers (see 3.2.1.). It should be noted that high affinity glutamate transporters have not yet been located at synapses of climbing fibers in immunocytochemical studies with antibodies against high-affinity glutamate transporters, although this possibility is still open since a detailed electron microscopical analysis of the cerebellar molecular layer has not yet been done (Rothstein et al., 1994). (4) Physiological studies suggested that the distal region of the Purkinje cell dendrites was relatively less sensitive towards aspartate as compared to glutamate than the proximal dendrites (Cr6pel et al., 1982). Since climbing fibers chiefly innervate the proximal two-thirds of the Purkinje cell dendritic tree (Palay and Chan-Palay, 1974), these data would be consistent with the proposal that aspartate is a climbing fiber transmitter, whereas glutamate is the transmitter of the parallel fibers (see Cu6nod et al., 1989). Voltage-clamp studies of Purkinje cells in slices, however, suggest that climbing fibers 55

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Fig. 43. Photomicrographs of consecutive semithin sections from rat inferior olive stained with antisera to aspartate, glutamate and GABA, respectively. All neurons (arrows) in this field are labelled for aspartate and glutamate, but unlabelled for GABA. Glial cells (arrowheads; identity established on the basis of electron microscopic analysis of adjacent sections) contain little or no GABA and glutamate immunoreactivities, but are moderately stained with the aspartate antiserum. Asterisks indicate fiber bundles. Scale bar = 50 r Insets show test sections incubated together with the respective tissue sections. The test antigens are GABA (1), glutamate (2), taurine (3), glycine (4), 'none' (5), aspartate (6), and glutamate (7). Note selective staining of the respective amino acid conjugates. Zhang et al. (1990). (

and parallel fibers activate the same type of glutamate receptors (Llano et al., 1991). Summarizing, one may conclude that the case for aspartate as the principal neurotransmitter of climbing fibers is far from being conclusive. Zhang et al. (1990), who compared glutamate- and aspartate-like immunoreactivities in the neurons of the inferior olive and climbing fibers in rat and baboon (Papaio anubis), showed that glutamate and aspartate-like immunoreactivities were co-localized in all neurons of the inferior olive, with a slightly heavier staining in the principal olive (Fig. 43). Significant glutamate-like, but little aspartate-like labelling, however, was recognized over climbing fiber profiles and, therefore, it was concluded that glutamate and not aspartate is the most likely transmitter of the climbing fibers (see also Zhang and Ottersen, 1993). It was also concluded that the presence of aspartate-like immunoreactivity in cell bodies is an unreliable indicator of transmitter identity.

Homocysteate Cu6nod et al. (1989) reported on the results of a series of experiments on K+-induced release of different transmitters by the cerebellum of the rat, after previous destruction of the inferior olive by 3-acetylpyridine. Release of aspartate was found to be decreased compared to the controls, with the main decrease occurring in the hemisphere. Values for the vermis were only slightly lower than in normal rats. This difference might be explained by a relative sparing of neurons in the caudal inferior olive, that project to the vermis. Decreased values after 3-acetylpyridine treatment were also found for adenosine (see Section 3.5) and for homocysteic acid. For the release of these substances no differences were noticed between vermis and hemisphere. Homocysteic acid was originally considered as a transmitter of the climbing fibers (Grandes et al., 1989), but proved to be located in Bergmann glia (Figs 44 and 45) (Cu6nod et al., 1990; Grandes et al., 1991). Climbing fibers, therefore, interact with Bergmann glia, both in the release of homocysteic acid and in 5'-nucleotidase-regulated adenosine release (see Section 3.5). Immunocytochemical studies have shown that subpopulations of climbing fibers may use peptides as a neurotransmitter, including somatostatine, corticotrophin-releasing factor and enkephalin. Their distribution and characteristics will be discussed in Section 6.3.4.

3.2.3. Granule cells and parallel fibers In early studies it was found that in 'staggerer', 'weaver' and 'reeler' mutant mice which have almost complete or partial loss of their granule cells (McBride et al., 1976a; Hudson et al., 1976) and in rats or mice that lost their granule cells by a viral infection or 57

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postnatal X-irradiation (McBride et al., 1976b; Rohde et al., 1979), glutamate was depleted. However, the interpretation of this finding is not immediately clear, because mossy fiber terminals and inhibitory interneurons of the cerebellar cortex also may have been affected. Furthermore, it proved difficult to exclude aspartate as a transmitter of granule cells (Rohde et al., 1979; Roffler-Tarlov and Turey, 1982). Also the demonstration of Garthwaite and Garthwaite (1985) that granule cells in slices accumulate [3H]Daspartate did not provide conclusive evidence about the nature of the neurotransmitter used by parallel fibers. Immunocytochemical studies strongly support glutamate as the neurotransmitter of the parallel fibers. Thus, glutamate-like immunoreactivity but no other amino acids were enriched over parallel fiber terminals in rat (Ottersen and Storm-Mathisen, 1984a,b, 1987; Ottersen et al., 1987, 1990; Liu et al., 1989) (Fig. 40), cat (Somogyi et al., 1986) and monkey (Zhang et al., 1990). Also electrophysiological experiments are in favour of glutamate as the neurotransmitter at the parallel fiber-Purkinje cell synapse (Barbour, 1993 and references therein).

~

[!2s .

Fig. 44. Immunocytochemical localization of homocysteate (HCA) in Bergmann glia with polyclonal antiHCA antibodies. A. Test system mimicking immunocytochemical procedure. Conjugates are assembled in 'sandwich' construction with tissue as spacer and contain the following compounds (from top to bottom): HCA, Glu (glutamate), Asp (aspartate), Tau (taurine), Gly (glycine), GABA (~,-aminobutyric acid), L-Ala (L-alanine), fl-Ala (fl-alanine), Htau (homotaurine), Hypotau (hypotaurine), Gline (glutamine), Ca (cysteate), CSA (cysteine sulphinate), HCSA (homocysteine sulphinate), Cys (cysteine), Cyt (cystine), Met (methionine), carnosine, Hcys (homocysteine), cystathionine, gluta-thione, homocarnosine, y-Glu-Glu (y-glutamyl glutamate), fl-L-Asp-Gly (fl-L-aspartyl glycine), no AA (no amino acid conjugated to glutaraldehyde-treated rat brain protein). B. Pattern of HCA-like immunoreactivity in low-power view of rat cerebellar cortex in semithin section. Double arrow: fibrous, radially oriented immunoreactive element. Arrowheads, stained varicosities in association with Purkinje cell dendrites. C. Pattern of HCA-like immunoreactivity in rat cerebellar section pretreated with 3-acetylpyridine 10 days previously and degeneration of the inferior olive. No changes in the distribution of HCA are apparent. Bars 50/lm. Grandes et al. (1991).

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Fig. 45. Immunocytochemical localization of homocysteate (HCA) with polyclonal anti-HCA antibodies. A. Staining pattern in section close to that in Fig. 44B at higher magnification. Cell (asterisk) and capillary (circle) used as landmarks in A and B. B and C. Electron micrographs from ultrathin section immediately preceding semithin section in A. The HCA-immunoreactive varicosities indicated with arrows in (A) were identified as parts of the glial sheath surrounding Purkinje cell dendrites (d) in B and C. Bars: 10/lm in A, 5/lm in B, 1 j~m in C. Grandes et al. (1991).

Specific markers for granule cells are few. Seyfried et al. (1983), concluded from biochemical analysis in 'weaver' mutant mice that the ganglioside GDIA was more concentrated in granule cells. Webb and Woodhams (1984) developed three monoclonal 59

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antibodies (G-l-3; 7-8D2 and 8-20-1), that recognize cell surface antigens expressed by rat granule cells and their axons (see also Reynolds and Wilkin, 1988). Calcium-binding proteins, with the exception of calretinin (Rogers, 1989; Arai et al., 1991; Kadowaki et al., 1993; Floris et al., 1994) have not been localized in granule cells. Proteine kinase C (PKC) e, flI and II, e and ~"are expressed by rat granule cells (Ase et al., 1988; Wetsel et al., 1992; Chen and Hillman, 1993) (Table 1). 3.3. LOCALIZATION OF GLUTAMATE RECEPTORS

3.3.1. lonotropic glutamate receptors Glutamate activates two main classes of glutamate receptors, the ionotropic and metabotropic glutamate receptors. The ionotropic receptors are receptor/channel complexes that can be categorized into three groups according to their differential sensitivity to agonist ligands, as ~-amino-3-hydroxy-5-methyl-4-isoxazole proprionic acid (AMPA) receptors, formerly known as quisqualate receptors, kainate receptors, and N-methylD-aspartate (NMDA) receptors (Monaghan et al., 1989; Mayer and Miller, 1990; Westbrook, 1994). The non-NMDA (AMPA and kainate) receptors display rapid kinetics. They are typically inhibited by 7-cyano-7-nitroquinoxaline-2,3-dione (CNQX), are permeable to monovalent cations (Na+, K+), but mostly impermeant to Ca 2+, and have been implicated in fast excitatory synaptic transmission (Mayer and Westbrook, 1987; Jahr and Lester, 1992). NMDA receptor channels, instead, have relatively slow kinetics, are also permeable to Ca 2+ ions, and are typically inhibited by D-2-amino-5-phosphonovalerate (APV). NMDA receptors are characterized by a voltage-dependent channel block by MgZ+-ions. They are dependent on, and are equipped with a coagonist site for glycine. Apart from their role in excitatory synaptic transmission, NMDA receptors have been implicated in synaptic plasticity and in developmental processes like cell migration and synaps formation (Collingridge and Singer, 1990). AMPA receptors

AMPA receptors have been autoradiographically labelled with [3H]AMPA and the antagonist [3H]CNQX: [3H]AMPA binding is moderately high over the rodent (Rainbow et al., 1984b; Monaghan et al., 1984; Nielsen et al., 1990; Garcia-Ladona et al., 1991; Makowiec et al., 1991) and human (Jansen et al., 1990) cerebellum, and is higher over the molecular than over the granular layer. [3H]CNQX binding sites are preferentially localized over the molecular layer, but cerebellar [3H]CNQX binding is relatively higher than [3H]AMPA binding, when the two are compared to binding levels of both ligands in other brain areas (e.g. see Fig. 6 in Nielsen et al., 1990). This difference is not due to [3H]CNQX binding to kainate receptors since these receptors are preferentially localized in the granular layer. Both [3H]AMPA and [3H]CNQX binding in the molecular layer was decreased in Purkinje cell deficient (pcd) mutant mice, but strongly upregulated in granuloprival mice (Makowiec et al., 1991). These observations favour a primary localization of AMPA receptors on Purkinje cells and an upregulation of the number of AMPA receptors on Purkinje cells as a consequence of deafferentation (Makowiec et al., 1991). Originally, AMPA receptors were assessed as quisqualate-sensitive [3H]glutamate binding sites (Cha et al., 1988, and references therein). Quisqualate-sensitive [3H]glutamate binding is strongly increased by the presence of CaC12, and is relatively high in the cerebellar molecular layer. CaC12-dependent quisqualate-sensitive [3H]glutamate bind60

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ing over the molecular layer, however, is largely insensitive to AMPA (Cha et al., 1988). These sites most likely correspond to the quisqualate-sensitive metabotropic glutamate receptors (Young et al., 1991), that have been recently demonstrated to be expressed at high levels by Purkinje cells (see Section 3.3.2.). Kainate receptors

High-affinity [3H]kainate binding sites predominate in the granular layer in rat (Monaghan and Cotman, 1982; Olson et al., 1987; Cambray-Deakin et al., 1990; Bahn et al., 1994) and man (Jansen et al., 1990). Low to moderate levels of [3H]kainate binding occur in the rat cerebellar nuclei. [3H]Kainate binding is not affected in Purkinje cell deficient (pcd) or 'nervous' mutant mice, but is decreased in granuloprival mice (Griesser et al., 1982). This decrease concerns the granular but not the molecular layer (Olson, 1987; Makowiec et al., 1991). Henke et al. (1981) noted a high level of low-affinity [3H]kainate binding sites in the molecular layer of pigeon cerebellum. Similar [3H]kainate binding sites were also labelled in the chicken cerebellum (Henley and Barnard, 1990), in fish (Maler and Monaghan, 1991) and in amphibian cerebellum, although in the amphibian kainate-binding sites seems to have somewhat different pharmacological and functional properties (reviewed in Henley, 1994). The chicken kainate binding sites could also be labelled by [3H]CNQX (Henley and Barnard, 1990). Several non-mammalian vertebrate kainate-binding proteins have been purified and cloned. These proteins display some homology towards mammalian ionotropic AMPA and kainate receptor subunits (see below), but are smaller (40-50 kDa instead of 100 kDa), and do not form functional receptors channels (reviewed by Hollman and Heinemann, 1994; Henley, 1994). In situ hybridisation and immunocytochemistry has shown that avian kainate-binding protein is localized in Bergmann glia (Fig. 95) (Somogyi et al., 1990; Gregor et al., 1992 and others). Somogyi et al. (1990) showed that immunostaining with a monoclonal antibody (IX-50) against chicken kainate-binding protein, was also localized in Bergmann glia in the cerebellum of fish. Frog kainatebinding protein, however, is widely distributed throughout the frog brain. High receptor densities were found in cerebellum, but their cellular distribution has not yet been reported (Dechesne et al., 1990; Wenthold et al., 1990). N M D A receptors

The distribution of NMDA receptors has been autoradiographically determined as NMDA-replaceable [3H]glutamate binding sites. In rat (Greenamyre et al., 1985; Monaghan and Cotman, 1985) and human cerebellum (Jansen et al., 1990), moderate densities of binding sites are found over the granular layer and in the cerebellar nuclei, whereas binding over the molecular layer is low. Olson et al. (1987) and Makowiec et al. (1991) reported that NMDA-sensitive [3H]glutamate binding is unchanged in Purkinje cell deficient (pcd) mutant mice, but that the density of binding sites is considerable reduced over the granular layer in granuloprival mice. These data suggest that NMDAbinding sites are absent on Purkinje cell dendrites and, instead, are present on granule cells and perhaps on stellate, basket and Golgi cells. Using different ligands including the competitive antagonist [3H]-2-carboxypiperazine-4-yl-propyl-l-phosphonic acid ([3H]CPP), [3H]glycine that specifically binds to the glycine coagonist site of NMDA receptors, and the non-competetive channel blockers [3H]MKS01 and [3H]-N-[1-(2-thienyl)cyclohexyl]-3,4-piperidine ([3H]TCP), it was found that the pharmacological properties of NMDA receptors in the cerebellar 61

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cortex were different from those in other brain areas (reviewed in Monaghan and Anderson, 1991). Notably, cerebellar NMDA receptors label poorly with the noncompetetive channel blockers [3H]MK801 and [3H]TCP (Maragos et al., 1988; Monaghan and Anderson, 1991). Monaghan and coworkers recognized at least 4 pharmacologically distinct NMDA receptor types throughout the brain and recently demonstrated that their pharmacological heterogeneity reflects differences in subunit composition (see below; Buller et al., 1994). They identified two populations of NMDA receptors in the cerebellar cortex. One population of 'antagonist-prefering' sites, that can be labelled by [3H]CPP, is present throughout the brain, and represents NMDA receptors containing NR2A subunits. The second population consists of the 'cerebellar-like' sites, where competitive antagonists and the agonist homoquinolinate are relatively ineffective in displacing the NMDA-sensitive [3H]glutamate binding. They reflect the presence of NR2C subunit, that is uniquely expressed by cerebellar granule cells (Buller et al., 1994). The low level of [3H]MK801 and [3H]TCP binding in the cerebellum remains to be explained. Distribution of subunits

Like other classes of ionotropic receptors functional glutamate receptor channel complexes are multimeric proteins. Recent molecular cloning studies have revealed families of AMPA (GluR1-GluR4, also named GluRA-GluRD), kainate (GluR5-GluR7, and KA1 and KA2), NMDA (NR1, named ~'1 in mice, and NR2A-NR2D, named el-e4 in mice) and orphan (~1 and ~2) glutamate receptor subunits (reviewed in Nakanishi, 1992; Sommer and Seeburg, 1992; Hollman and Heinemann, 1994). The diversity of glutamate receptor subunits is further increased through alternative splicing that primarily involves the AMPA receptor subunits GluR1-GluR4, each of which exists in two versions, i.e. flip or flop, and the NR 1 subunit, that has eight splice variants. Combinatorial expression studies have demonstrated that the subunits aggregate into functional receptor channels in the homomeric as well as the heteromeric configuration. Thus multiple functionally distinct forms of each receptor type can be formed through different combinations of subunits (see below). In situ hybridisation (KeinS.nen et al., 1990; Monyer et al., 1991, 1994; Araki et al., 1993; Sato et al., 1993; Wisden and Seeburg, 1993; Akazawa et al., 1994; Laurie and Seeburg, 1994; Watanabe et al., 1994; and others) and immunocytochemical studies with antibodies for specific subunits (Martin et al., 1992, 1993; Petralia and Wenthold, 1992; Brose et al., 1993; Baude et al., 1994; Nusser et al., 1994; Petralia et al., 1994a,b,c; Jaarsma et al., 1995b) have shown that subunits are heterogeneously distributed throughout the cerebellum, each cell type expressing a characteristic set of subunits (see Table 2). AMPA subunits

AMPA receptor subunits are not only expressed by cerebellar neurons, but also by Bergmann glia, that express high levels of GluR 1 (GluRA) and GluR4 (GluRD) mRNA (Table 2, Fig. 46). GluR1 subunit mRNA is also expressed by Purkinje cells but not by other cerebellar cells (Kein~inen et al., 1990; Monyer et al., 1991; Sato et al., 1993). GluR4 mRNA in addition to Bergmann glial cells, is produced by granule cells and neurons of the deep nuclei. Granule cells express a GluR4 splice variant exclusively found in the cerebellum, GluR4c, consisting of GluR4 with the flop module and a truncated C-terminus (Gallo et al., 1992). GluR2 (GluRB) mRNA is found over the granular and molecular layers, in Purkinje cells, and in cells of the deep nuclei (Fig. 46). 62

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GluR3 (GluRC) mRNA is not expressed in granule cells, but occurs in Golgi cells, stellate and basket cells, Purkinje cells and cells in the deep nuclei (Fig. 46) (KeinS.nen et al., 1990; Monyer et al., 1991; Sato et al., 1993). The distribution of AMPA subunits has been studied immunocytochemically with antibodies specific for GluR1, GluR2/3, and GluR4 (Martin et al., 1992; Petralia and Wenthold, 1992; Baude et al., 1994; Nusser et al., 1994; reviewed in Jaarsma et al., 1995b). The processes of the Bergmann glia are densely immunostained for GluR1 and GluR4 (Fig. 47) confirming in situ hybridisation. Electron microscopy showed that GluR1 immunoreactivity was localized throughout the cytoplasma membrane (Baude et al., 1994). Dense immunostaining was associated with the processes of Bergmann fibers ensheathing PC spines and the attached synaptic varicosities of parallel fibers and climbing fibers (Fig. 48). This indicates that AMPA receptors on Bergmann glia may be activated by glutamate released by parallel fibers or climbing fibers, allthough there is no clue as yet of the functional role of glial cell activation (see discussion Baude et al., 1994, but also Mfiller et al., 1992). Purkinje cells are weakly-to-moderately immunopositive for GluR1. Dense GluR1 immunolabelling was found at the post-synaptic membrane specialisations of the dendritic spines of Purkinje cells, facing parallel and climbing fiber boutons (Fig. 48). The post-synaptic membranes of the parallel fiber-Purkinje cell and the climbing fiberPurkinje cell synapses are also stongly immunoreactive for GluR2/3 (Nusser et al., 1994; Jaarsma et al., 1995b). The GluR2/3 antibodies immunoreact with all cerebellar neurons. The perikarya and dendritic arbors of Purkinje cells densely immunostain, whereas

TABLE Type

AMPA

Kainate

NMDA

orphan

2.

Distribution of glutamate receptor subunit mRNAs in rat cerebellum Subunit

Cell t y p e PC

GrC

GoC

GluR1

+ flip

-

-

-

+ + flip

-

GluR2

+ + flip/flop

+ flip

+

+

-

++

GluR3

+ flip

-

++

++

-

+

GluR4

-

+ 4c-flop

-

-

+ + flip

+

GluR5 GluR6

+ -

. ++

GluR7

-

-

-

+

KA1

+

.

.

BC/Stc

. .

Bg

.

.

.

.

-

.

+

.

.

DCN

.

.

KA2

-

++

-

-

-

+

NR1

+(NRI-b)

++(NRI-a)

+

+

-

++

NR2A

-

+

-

-

-

+

NR2B

.

NR2C

-

++

.

NR2D

-

-

+

-

+

delta 1

+

.

delta2

.

.

.

.

.

.

.

.

.

+

. .

.

. .

. .

. .

S y m b o l s : - , n o t d e t e c t e d ; +, p o s i t i v e ; + + , s t r o n g l y p o s i t i v e ; P C , P u r k i n j e cells; G r C , g r a n u l e cells; G o C , G o l g i cells; B C , b a s k e t cells; St, s t e l l a t e cells; Bg, B e r g m a n n

glia; D C N ,

deep cerebellar nuclei.

B a s e d o n d a t a f r o m K e i n ~ n e n et al., 1990; M o n y e r et al., 1991, 1994; L a m b o l e z et al., 1992; A r a k i et al., 1993; L o m e l i et al., 1992, 1993; S a t o et al., 1993; W i s d e n a n d S e e b u r g , 1993; A k a z a w a S e e b u r g , 1994; W a t a n a b e

et al., 1994; L a u r i e a n d

et al., 1994.

63

P

Fig. 46. In situ hybridization of AMPA glutamate receptor mRNAs in sections of rat cerebellum. A. GluRl (GluRA) mRNA distribution; arrow heads indicate continuous line of silver grains along the Purkinje-Begmann layer. B. GluR2 (GluRB) mRNA; arrow heads indicate labelled Purkinje cells. C. GluR3 (GluRC) mRNA; small arrow heads indicate clusters of silver grains in molecular layer over stellate-basket cells. D. GluR4 (GluRD); arrow heads as in (A). gr, granule cell layer; mol, molecular layer; p, Purkinje cells; wm, white matter. Scale bar 500 fim. Keinanen et al. (1990).

a

& h

The cerebellum." chemoarchitecture and anatomy

Ch. I

5',.2 uletl

Fig. 47. Sagittal sections of the rat cerebellar cortex immuno-labelled with antibodies to GluR1 (a), GluR2/3 (b,e), and GluR4 (c,d). As, astrocyte-like cells; BG, Bergmann glial processes; Go, Golgi cell; Gr, granular layer; L, Lugaro cell; Mo, molecular layer; Pj, Purkinje cell body; WM, white matter; small arrow, Purkinje cell dendrite; asterisks, Bergmann glial cell body; arrow head, basket/stellate cell. Petralia and Wenthold (1992).

light-to-moderate staining neurons occur in basket/stellate cells, Golgi cells and granule cells (Fig. 47) (Martin et al., 1992, 1993; Petralia and Wenthold, 1992; Jaarsma et al., 1995b). Unipolar brush cells are also strongly GluR2/3-immunopositive (Jaarsma et al., 1995b). Dense and moderate GluR2/3-staining was found in the perikarya and neuropil of the deep nuclei, respectively. Using electronmicroscopic immunogold protocols, that allow precise ultrastructural localization of the immunoreaction product, Nusser et al. (1994) obtained stong proof that GluR2/3 immunoreactivity is associated with postsynaptic membrane specialisations of excitatory synapses in the cerebellar cortex (Fig. 49B, C, F). Their data indicate that GluR2/3 immunoreactivity is considerably stronger at parallel fiber-Purkinje cell, climbing fiber-Purkinje cell and parallel fiber-stellate cell synapses than at mossy fiber-granule cell synapses (compare Figs 49B and C with F). Conventional peroxidase-DAB (3,3'-diaminobenzidine tetrahydrochloride)-immuno65

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J. Voogd, D. Jaarsma and E. Marani

electron microscopy also indicates that GluR2/3 immunoreactivity is relatively weak at the mossy fiber-granule synapses (Jaarsma et al., 1995b). The post-synaptic membranes of the giant mossy fiber-unipolar brush cell synapses are, however, strongly GluR2/3 immunopositive (Jaarsma et al., 1995b). The GluR4 antibodies, in addition to the Bergmann glia, moderately immunostain the granular layer, and the neuropil and perikarya in the the deep nuclei (Fig. 47). It was originally assumed that GluR4-immunostaining in the granular layer was associated with granule cells (Martin et al., 1992, 1993; Petralia and Wenthold, 1992), but electron microscopy showed that GluR4 immunoreactivity is localized in the astroglia (Jaarsma et al., 1995b). Thus granular layer astroglia like Bergmann glia express AMPA receptor subunits, but unlike the Bergmann glia, do not have GluR1. The absence of GluR4-immunoreactivity in granule cells can be explained by the fact that granule cells primarily express an atypical form of GluR4, GluR4c (see above), that is not recognized by the GluR4 antibodies currently available. AMPA receptors are believed to mediate most of the fast excitatory neurotransmission in the brain, including the cerebellum. Concordantly the types of AMPA receptor subunits expressed by a cell largely determine the characteristics of the fast excitatory responses (see Jonas and Spruston, 1994). The GluR2 subunit dominate the AMPA receptor channel behavior, in that homomeric GluR2 channels as well as heteromeric

Fig. 48. Electron micrograph of the synaptic distribution of immunoreactivity for the GluR 1 subunit of the AMPA receptor in rat cerebellum as detected by an antibody against the carboxy-terminal (intracellular) region of GluR1. A. A spine (s) emerging from a Purkinje cell dendrite (Pd) establishes an immunopositive type 1 synapse (solid arrows) with a parallel fiber terminal (pft). Intra-cellular immunoreactivity is present inside Bergmann glial cell processes along dendritic elements (e.g., open arrow). B. The peroxidase reaction end-product labels the postsynaptic density (psd) at the intracellular face of the postsynaptic membrane (pom) and not the synaptic cleft between the presyaptic (pem) and postsynaptic (pom) membranes. Scale bars in A = 0.5 r in B = 0.1 r Baude et al. (1994).

66

The cerebellum." chemoarchitecture and anatomy

Ch. I

channels formed with the participation of GluR2 show the properties of 'typical' AMPA receptors, i.e. linear current-voltage relations and a relative impermeability to Ca 2+. Homo- or heteromeric channels without GluR2, instead, display inward rectification and are permeable to Ca 2+ and other divalent cations (for references see Sommer and Seeburg, 1992; and Hollman and Heinemann, 1994). These channels usually are not found in neuronal cells, concordant with the notion that most neuronal cells express GluR2, but are present in Bergmann glial cells (Mfiller et al., 1992; but see also Burnashev et al., 1992). In accordance with the presence of high levels of GluR 1 and GluR4 but absence of GluR2 in these cells. By combining two powerful methods, i.e. the patch-clamp technique to characterize the properties of native receptor channels in single cells in brain slices, followed by single cell PCR-amplification methods to analyse the mRNA contents of the respective cells semiquantitatively, Jonas et al. (1994) recently showed that the CaZ+-permeability of native AMPA receptor channels in cerebral cortical cells is related to the relative abundance of GluR2 subunit mRNA in the respective cells. Thus inhibitory interneurons of the cerebral cortex have low GluR2/non-GluR2 ratios (-- 0.3) and highly Ca 2+permeable AMPA receptors (which, however, display linear current-voltage relations unlike 'Bergmann-glial' AMPA receptors), whereas pyramidal cells, which have a relatively high GluR2/non-GluR2 mRNA ratio (-- 3), contain CaZ+-impermeable AMPA receptors. PCR-amplification analysis of AMPA subunit m R N A of Purkinje cells, indicates that GluR2 mRNAs are more abundant than GluR1 and GluR3 mRNAs (Lambolez et al., 1992), implying that Purkinje cells express weakly CaZ+-permeable AMPA receptors. Also in (pooled) granule cells GluR2 mRNA is more abundant than non-GluR2 (GluR4) mRNA. PCR-amplification analysis has not yet been done for other cerebellar cells. One might speculate that the basket, stellate and Golgi cells express CaZ+-permeable AMPA receptor like the inhibitory interneurons of the cerebral cortex, since according to in situ hybridisation data they seem to express relative high levels of GIuR3 compared to GluR2 (see Table 2). AMPA receptors made from different subunits may have different desensitization kinetics. Desensitization is particularly fast for AMPA receptors formed with GluR3flop or GluR4-flop (Mosbacher et al., 1994). Granule cells produce GluR4-flop and GluR2 (Table 2), and therefore are likely to have fast (submillisecond) desensitizing AMPA receptors. This could explain the very fast decay kinetics of non-NMDA component of the excitatory post-synaptic currents (EPSCs) at the mossy fiber-granule cell synapses (Silver et al., 1992; Rossi et al., 1995). If this holds true this would imply that the length of the excitatory responses at the mossy fiber-granule cell synapses is largely controlled by the desensitization properties of the AMPA receptors and does not depend upon the time course of transmitter removal, that may be relatively slow at these synapses (Jonas and Spruston, 1994) (see Section 3.2.1.). Purkinje cells express GluRl-flip, GluR2-flip and -flop, and GluR3-flip mRNA (Lambolez et al., 1992) that form AMPA receptors with desensitization time constants 3-5 times slower than 'GluR4-flop-GluR2' channels (Mosbacher et al., 1994). Concordantly, AMPA receptors in Purkinje cells appear to have relatively slow desensitization kinetics (Barbour et al., 1994). Also the decay phases of AMPA receptor-mediated EPSCs in Purkinje cells after parallel fiber or climbing fiber activation, have slow time constants (Perkel et al., 1990; Llano et al., 1991; Barbour et al., 1994). Interestingly stellate/basket cells, that express AMPA receptors with the same desensitization kinetics as Purkinje cell AMPA receptors, showed much faster decaying parallel fibers EPSCs (Barbour et al., 1994). Barbour et al. (1994) concluded that glutamate is rapidly cleared 67

Ch. I

J. Voogd, D. Jaarsma and E. Marani

at the parallel fiber-stellate/basket cell synapses, resulting in rapid deactivation of postsynaptic AMPA receptors, whereas synaptically released glutamate seems to be present during a prolonged time at Purkinje cell synapses. According to H/~usser (1994) EPSCs of climbing fiber-Purkinje cell synapses have decay time constants that are slower than parallel fiber-evoked EPSCs in Purkinje cells, which may be explained by the fact that clearance of glutamate at climbing fiber synapses is slower due to their larger size.

68

The cerebellum." chemoarchitecture and anatomy

Ch. I

Fig. 49. Electron micrographs showing the subsynaptic segregation of GluR2/3 AMPA receptor subunits (GluR B/C/4c) and the metabotropic mGluRl~ glutamate receptor (mGluR1; see Section 3.3.2) as revealed by post-embedding immunogold labelling. A and B. Consecutive sections of the same synaptic junctions showing that immunoparticles for mGluRl~ (double arrows in A) are concentrated at the edge, whereas immunoparticles for GluR2/3 (arrows in B) are concentrated in the main body of synaptic junctions established by parallel (pft) and climbing (cft) fiber terminals with spines (s) of Purkinje cell dendrites (Pd). Note that mGluRlcz is often localized extrasynaptically (double arrow heads in A). C. Immunoreactivity for GluR2/3 (arrows) was always very strong on basket and stellate (Stc) cells. D and E. Double immunolabelling of mGluRl~ (large particles, double arrows) and GluR2/3 (small particles, arrows) immunoreactivity in the synapses on spines (s) of Purkinje cells, confirming synaptic segregated subsynaptic localization of mGluR1 and GluR2/3. The synapse in E is from Triton treated material. F. Generally a lower density of immunoparticles for GluR2/3 (arrows) has been found in synapses between mossy fiber terminals (mt) and granule cell dendrites (d) than in the parallel fiber synapses (compare to B and C). Scale bars = 0.1 r in A,B,D,E, 0.2 r in C,F. Nusser et al. (1994). (

Kainate subunits

Kainate receptor subunits are most prominent in granule cells, which express high amounts of both GluR6 and KA2 mRNA (Fig. 50) (Wisden and Seeburg, 1993). Purkinje cells express moderate levels of GluR5 and low levels of KA1 mRNAs; basket and stellate cells express GluR7 mRNA, and neurons of the deep cerebellar nuclei produce GluR7 and KA2 mRNAs (Table 2, Fig. 50) (Wisden and Seeburg, 1993). The high level of kainate receptor mRNA expression by granule cells is in accordance with the preferential binding of [3H]kainate over the granular layer (see above, but also Bahn et al., 1994). Immunocytochemical studies with antibodies specific for GluR6 and GluR7 show that dense GluR6/7-immunostaining occurred over the granule cell layer, where it was associated with granule cell perikarya and dendrites (Petralia et al., 1994c; Jaarsma et al., 1995b). The post-synaptic membranes of the mossy fiber-granule cell synapses were strongly immunoreactive for GluR6/7 (Jaarsma et al., 1995b). Stellate and basket cells and cells in the deep cerebellar nuclei were also immunoreactive for GluR6/7. KA2 immunoreactivity was relatively low in the cerebellar cortex and was concentrated over the glomeruli and neurons in the deep nuclei (Petralia et al., 1994c; Jaarsma et al., 1995b). Recombinant expression studies have shown that GluR5 and GluR6 form glutamategated channels in the homomeric configuration as well as in the heteromeric configuration with KA1 and KA2, whereas KA1 and KA2 do not assemble into functional receptor channel complexes (reviewed in Wisden and Seeburg, 1993). Thus functional kainate receptors can be formed in several cerebellar cells, in particular in granule cells expressing significant levels of GluR6 and KA2. Unequivocal physiological evidence for the presence of kainate receptors in the cerebellum (as well as in other brain areas) is, however, still lacking (see discussion Wisden and Seeburg, 1993). It should be noted that kainate in spite of its low affinity for AMPA receptors, potently activates AMPA receptors, and that excitatory responses evoked by kainate in brain tissue are generally mediated through AMPA receptors. Recombinant kainate receptors have been demonstated to desensitize very rapidly, which in part may explain why kainate receptor responses have not been detected (Wisden and Seeburg, 1993). Another possibility is that kainate receptor responses are masked by AMPA receptors, which are assumed to be present in much higher concentrations in neurons (Wenthold et al., 1994 and references therein). Autoradiographic (see above) and immunocytochemical studies (Jaarsma et al., 1995b), however, suggest that kainate receptors predominate over AMPA recep69

Ch. I

~LuR-5

J. Voogd, D. Jaarsma and E. Marani

,~ .~LuR ::6

B .3LuR 7

C Mol

Gr P "If ;:' l"'.s,"''~,;,"\ T (

'

i

f

Po

Fig. 50. Distribution of GluR5 (A), GluR6 (B), GluR7 (C), KA-1 (D), KA-2 (E) of subunits in RNAs of high-affinity kainate receptor mRNAs in coronal sections at level of the cerebellum of the rat. Gr, granular layer; LC, locus coeruleus; Mol, molecular layer; P, Purkinje cell layer; Po, pontine nuclei. Scale bars: 2.3 mm. Wisden and Seeburg (1993).

tors in granule cells, and may significantly contribute to excitatory neurotransmission at the mossy fiber-granule cell synapses. N M D A subunits Functional NMDA receptors are believed to be generated as heteromeric assemblies of NR1 subunits with members of the NR2 subunit family. The pharmacological and kinetic heterogeneity of NMDA receptors seems to be primarily dependent upon the type of NR2 subunit (Monyer et al., 1992; Meguro et al., 1992; Nakanishi, 1992; Buller et al., 1994), although NR1 diversity generated through alternative splicing may also contribute to NMDA receptor heterogeneity (Buller et al., 1994; Hollman and Heineman, 1994). Essentially all cerebellar neurons seem to express significant levels of NR1 mRNA (Table 2, Fig. 51E) (Moriyoshi et al., 1991). The main splice variants produced in the cerebellum are NR 1-2 (with 3'-end deletion 1) and to a lesser extent NR1-4 (with 3'-end deletions 1 and 2; Laurie and Seeburg, 1994). There is a remarkable difference between the Purkinje cells and the other cells of the cerebellar cortex, in that Purkinje cells express high levels of the NRI-a forms (without 5'-insertion), whereas in the other cells the N 1-b splice variants (with 5'-insertion) predominate (Laurie and Seeburg, 1994). NR2 subunit mRNAs are heterogeneously expressed throughout cerebellar neurons (Table 2, Fig. 51) and show pronounced changes during development (Akazawa et al., 1994; Monyer et al., 1994; Watanabe et al., 1994). NR2 subunit mRNAs are most prominent in granule cells, that express high levels of NR2C mRNA and moderate levels of NR2A mRNAs in adult rodent cerebellum (Fig. 51). Interestingly, whereas NR2A mRNA expression in rodent granule cells begins early postnatally, NR2C first appears in later stages (postnatal day 10-11 in rat) in post-migratory cells of the internal granular layer. It apparently replaces NR2B, which is transiently expressed by cerebellar granule cells (Akazawa et al., 1994; Monyer et al., 1994; Watanabe et al., 1994). The expression of NR2C starts in granule cells of the caudal vermis (lobules VIII-X) and subsequently extends throughout the whole cerebellar cortex by postnatal day 13 (see Fig. 3N and O in Watanabe et al., 1994 and Fig. 7 in Akazawa et al., 1994). This pattern is compat-

70

The cerebellum." chemoarchitecture and anatomy

Ch. I

ible with the sequence of maturation of the granule cells (Altman, 1972). According to Akazawa et al. (1994) and Watanabe et al. (1994), but not Monyer et al. (1994), NR2C mRNA is also expressed in the external granular layer during the first postnatal days. NMDA receptors have been demonstrated to be critically involved in granule cell migration (Komuro and Rakic, 1993; Rossi et al., 1993). Since NR2B is transiently expressed by granule cells during the period of migration, one may speculate that receptors made with NR2B and NR1 (and possibly NR2A) may act as 'migration receptors'. The presence of multiple NMDA receptors in granule cells is consistent with the presence of multiple NR2 subunits and has recently been demonstrated with patchclamp methods (Farrant et al., 1994): Pre-migratory and migratory granule cells were shown to express NMDA receptor channels with conductancy properties of recombinant NMDA receptors formed by co-expression of NR 1 and NR2A or NR2B. Mature post-migratory cells, in addition, express 'low-conductance' NMDA receptor channels, which have the properties of NMDA receptors with NR2C (Monyer et al., 1994). Basket, stellate cells, Golgi cells and neurons in the cerebellar nuclei express NR2D mRNAs (Akazawa et al., 1994; Monyer et al., 1994; Watanabe et al., 1994). Neurons of the cerebellar nuclei also produce NR2A mRNA, but it is not clear whether NR2A and NR2D producing cells reflect distinct neuronal populations. It should be noted that NMDA receptors composed of NR1 and NR2D have very slow deactivation kinetics (roll = 4.8 s) (Monyer et al., 1994) and, therefore, may modulate the cell activity during many seconds even when the receptor channel has been briefly activated by glutamate (see discussion Monyer et al., 1994). Quinlan and Davies (1985) have provided indirect physiological evidence for the presence of NMDA receptors in stellate and basket cells, by showing that NMDA may induce inhibition of Purkinje cells. Also neurons of the deep cerebellar nuclei have been shown to display prominent NMDA responses in cerebellar slice cultures (Audinat et al., 1990).

,Sg~,'

.4

o ~

.._ ., /;.'_. . ;

,

.

_.

.

..

Fig. 51. Bright-field micrographs showing cellular distributions of the NMDA receptor channel subunit mRNAs in the cerebellar cortex of the adult mouse: (A) el (mouse homologue of NR2A) mRNA; (B) e2 (NR2B); (C) e3 (NR2C); (D) e4 (NR2D); and (E) ~'1 (NR1). Each photograph in the figure was taken from lobule V of the cerebellar vermis, and the expression patterns of the respective subunit mRNAs are identical to those in remaining regions of the cerebellum. Sections were counter-stained with toluidine blue. Arrows indicate cell bodies of the Purkinje cells. Gr, granular layer; Mol, molecular layer. Scale bar = 50 r Watanabe et al. (1994).

71

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J. Voogd, D. Jaarsma and E. Marani

The presence of NMDA receptors on Purkinje cells has been disputed. Some studies have supported the presence of NMDA-receptors on Purkinje cells (Sekiguchi et al., 1987), but in most studies no evidence of NMDA-receptors on Purkinje cells has been found (e.g. Audinat et al., 1990; Perkel et al., 1990; Llano et al., 1991; Farrant and Cull-Candy, 1991). Studies of Krupa and Cr6pel (1990) and Rosenmund et al. (1992) have indicated that NMDA receptors are present on most Purkinje cells during early post-natal life, but disappear with age. Both in situ hybridisation in rat and mouse and immunocytochemical studies in rat have shown that the NR1 subunit is expressed at high levels by Purkinje cells (Brose et al., 1993; Akazawa et al., 1994; Monyer et al., 1994; Petralia et al., 1994a; Watanabe et al., 1994). Petralia et al. (1994a) further demonstrated that NRl-immunoreactivity occur at the post-synaptic membrane specialisations in Purkinje cell spines. With respect to the NR2 subunits, Akazawa et al. (1994) found that rat Purkinje cells may express NR2D mRNA until post-natal day 8 and thereafter express low levels of NR2A mRNA. Accordingly Petralia et al. (1994b) observed that Purkinje cells display a low level of NR2A/B immunoreactivity, also in the post-synaptic densities of Purkinje cell dendritic spines, indicating that low levels of 'NR1-NR2A' receptors may be present at parallel fiber or climbing fiber synapses. Watanabe et al. (1994), however, found that mouse Purkinje cells only express low levels of NR2B (indicated as e2, which is the mouse homolog of NR2B) until one day postnatally, but not at any later stage, whereas according to Monyer et al. (1994) Purkinje cells do not produce any NR2 mRNA at any age. One may conclude from the in situ hybridisation and immunocytochemical data, that in spite of the presence of high levels of NR1 subunit, Purkinje cells both during development and in adulthood are likely to express none or only low amounts of functional NMDA receptors, which is in line with the aut0radiographic data. NR1 subunits can also form receptor-channel complexes in the homomeric configuration, but these channels produce very small currents and are, therefore, unlikely to contribute significantly to the excitatory actions of glutamate in Purkinje cells (Moriyoshi et al., 1991). Orphan receptors

51 and 52 are two related subunits isolated by homology screening. 51 is not produced in the rodent cerebellum, but 52 is selectively expressed by Purkinje cells (Araki et al., 1993; Lomeli et al., 1993). The subunit protein is distributed throughout the somatodendritic domain of the Purkinje Cells, similar to other glutamate receptor subunits (Araki et al., 1993). The function of S1 and 52 is not yet understood. The subunit protein does not bind glutamate receptor agonists and does not aggregate into functional receptors (Lomeli et al., 1993).

3.3.2. Metabotropic glutamate receptors Metabotropic glutamate receptors are coupled to G-proteins and modulate intracellular second messenger systems. The metabotropic glutamate receptors consist of at least seven subtypes that can be subdivided into three subgroups on the basis of sequence homology, agonist selectivity, and second messenger system (Nakanishi, 1992; Tanabe et al., 1992): (1) mGluR1 and mGluR5, that are coupled primarily to activation of phosphoinositide hydrolysis and are activated by quisqualate (QA) and 1S,3R-aminocyclopentane dicarboxylate (1S,3R-ACPD); (2) mGluR2 and mGluR3, that are coupled to inhibition of the cAMP cascade, are sensitive to pertussis toxin, and are activated by 72

The cerebellum." chemoarchitecture and anatomy

Ch. I

1S,3R-ACPD, but are insensitive to QA; and (3) mGluR4, mGluR6 and mGluR7, which are also coupled to inhibition of the cAMP cascade, and are potently activated by L-2-amino-4-phosphonobutyrate (L-AP4), but are insensitive to QA and 1S,3R-ACPD. Metabotropic glutamate receptors have been implicated in multiple neuronal processes including modulation of transmitter release, plasticity phenomena such as long term potentiation and long term depression, and other long term changes of neuronal functions (see Schoepp, 1994 for a review). With the exception of mGluR6 that is expressed only in retina, all metabotropic receptors are expressed in the cerebellum. mGluR1 mRNA is expressed to some extent by most cerebellar neurons (Shigemoto et al., 1992), but is found at very high levels in Purkinje cells. Immunocytochemistry shows that the mGluR1 protein is localized in the spines of Purkinje cell dendrites (Fig. 52) (Martin et al., 1992; Baude et al., 1993; Shigemoto et al., 1994). Dense mGluR1 immunostaining is also associated with the brushes of unipolar brush cells (Jaarsma, Mugnaini, Shigemoto et al., in preparation). Interestingly, mGluR1 immunostaining is not associated with the post-synaptic region of the giant mossy fiber-unipolar brush cell synapses, but instead, occurs at very high levels in spiny appendages and small branchlets that emanate from the dendritic stem that do not have synaptic specialisations (see Mugnaini et al., 1994). Recently workers from Somogyi's group (Baude et al., 1993; Nusser et al., 1994) demonstrated, with immunogold techniques, that mGluR1 immunoreactivity in Purkinje cell spines (as well as in other neurons) was never localized to the postsynaptic membrane specialisations of the synapses, but was associated with perisynaptic and extrasynaptic regions. This is in marked contrast with ionotropic glutamate receptor subunits that are primarily located at the postsynaptic membrane (Fig. 49) (Nusser et al., 1994). It has been proposed that, as a consequence of its peri-and extrasynaptic localization, mGluR1 is only activated during high frequency stimuli, because low frequency stimuli may not release enough glutamate to reach the perisynaptic receptors at significant concentrations (Baude et al., 1993; Nusser et al., 1994). It was originally reported by Kano and Kato (1987) that a QA/transAPCD-sensitive glutamate receptor is critically involved in the induction of long term depression (LTD) of parallel-fiber-Purkinje cell synapses, a cerebellar paradigm of synaptic plasticity that is induced following repetitive stimulation of parallel fibers in conjunction with climbing fiber input (Ito, 1989; Linden and Connor, 1993). Recently strong evidence has been obtained that mGluR1 plays a major role in cerebellar LTD: (1) the induction of LTD could be inhibited with antiserum that inactivated mGluR1 in an in vitro model of LTD (Shigemoto et al., 1994); and (2) LTD could not be induced in a mutant mouse lacking mGluR1 (Aiba et al., 1994). In these animals the anatomy of the cerebellum was not overtly disturbed. The Purkinje cells showed some minor morphological alterations, but had normal excitatory responses upon parallel fiber and climbing activation. Interestingly, the animals showed characteristic cerebellar symptoms such as ataxic gait and intention tremor, which suggest that mGluR1, possibly through its role in LTD, is important in cerebellar function. The mGluR5 receptor is selectively localized to a subpopulation of Golgi cells with the receptor protein localized throughout the somato-dendritic domain of the cells (Abe et al., 1992b; Shigemoto et al., 1993). Also mGluR2 and mGluR3 mRNA's are selectively expressed by Golgi cells (Ohishi et al., 1993, 1994), although mGluR3 may also occur in glial cells (Ohishi et al., 1994; Tanabe et al., 1993). Using an antibody selective for mGluR2 and mGluR3, Ohishi et al. (1994) found that mGluR2/3 immunoreactivity was strongest in Golgi axon terminals in the glomeruli (Figs 53 and 54). The Golgi axon terminals are not in close contact with mossy fibers, but the distance between mossy fiber 73

Ch. I

J. Voogd, D. Jaarsma and E. Marani

Fig. 52. A. Photomicrograph of semithin 3/lm thick plastic section of the nodulus of rat cerebellar cortex immunostained with an antibody against the carboxyterminus of the metabotropic glutamate receptor, mGluRl~z (antibody A52) (Shigemoto et al. 1994). Immunoreaction product in the molecular layer (ml) has a punctate distribution. Very little staining occur in the perikarya and primary dendrites of Purkinje cells (PC). In the granular layer moderate and dense immunoreactivity is localized to the perikarya and 'brushes' (open arrows) of unipolar brush cells (asterisks in cell nucleus), respectively. Bar = 20/lm. B. Electron micrograph of the molecular layer showing that puncta within the cerebellar molecular layer correspond to mGluRlctimmunoreactive spines (arrows) of Purkinje cells. Curved arrow point to an immunoreactive spine branching from an unlabelled Purkinje cell dendrite (PCd). pf, parallel fiber terminal. Bar = 0.5/lm. Courtesy of Jaarsma, Dino, Mugnaini, Ohishi and Shigemoto.

terminals and Golgi axon terminals is usually less than 1 ~tm, and it is possible that glutamate released from mossy fibers may diffuse into the intercellular space to activate mGluR2/3 on the Golgi cell axons (see Section 3.2.1.). mGluR2/3 in Golgi axon terminals may be involved in the regulation of inhibitory neurotransmitter release, which would imply that mossy fibers may directly influence inhibitory neurotransmission on granule cell dendrites (e.g. see discussion Ohishi et al., 1994). Both mGluR5 and mGluR2/3 antibodies immunostain subpopulations of Golgi cells (Shigemoto et al., 1993; Ohishi et al., 1994). mGluR2/3 immunoreactive Golgi cells constitute three-quarters of the total population of Golgi cells (defined as GABApositive, parvalbumin-negative cells of the granular layer), whereas only a small population of Golgi cells appears mGluR5 positive. Large mGluR2/3-positive Golgi cells were frequently encountered in the Purkinje cell layer and the superficial part of the granular layer (Fig. 53), and at least in part may represent the candelabrum cells as described by Lain6 and Axelrad (1994, see section 2). In contrast, large mGluR5-positive immunoreactive Golgi cells were mostly found deeper in the granular layer. This indicates that mGluR2/3 and mGluR5 positive Golgi cells represent different subpopulations of Golgi cells. It remains to be determined whether mGluR2/3 and mGluR5 positive cells are entirely exclusive or overlapping populations, and whether yet another 74

The cerebellum." chemoarchitecture and anatomy

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Fig. 53. a. Immunocytochemicallocalization of mGluR2/3 in a parasagittal section through the vermis of rat cerebellum. The most intense immunoreactivity is seen in the granular layer. Staining in the molecular layer is associated with Golgi cell dendrites. B. Drawing of the section shown in (a) showing presumed Golgi cell bodies with mGluR2/3 immunoreactivity (closed circles) and those without mGlu2/3 immunoreactivity (open circles). G, granular layer; M, molecular layer; P, Purkinje cell layer; W, white matter. Bar = 500 #m. Ohishi et al. (1994).

subpopulation exists that is both mGluR5 and mGluR2/3 negative. It is important to realize that mGluR2/3 as well as mGluR5 immunoreactivity is detected in both small and large Golgi cells, and that, therefore, the segregation of Golgi cells into mGluR2/3 respectively mGluR5-positive and negative cells does not correspond to previous classifications which were based on size (e.g see Palay and Chan-Palay, 1974; but also section 3.6.2.). The L-AP4-sensitive mGluR, m G l u R 4 is expressed at high levels by granule cells (Kristensen et al., 1993; Tanabe et al., 1993), whereas mGluR7 m R N A is produced by Purkinje cells (Okamoto et al., 1994; Saugstadt et al., 1994). Physiological data indicate that the L-AP4 sensitive mGluRs are predominantly located presynaptically, where they may act as autoreceptors to regulate glutamate release. Studies in turtle suggest the presence of a L-AP4-sensitive presynaptic glutamate receptor at the parallel fiber-Purkinje cell synapse (Larson-Prior et al., 1989). Thus, possibly, mGluR4 is located presynaptically on parallel fiber boutons. The ultrastructural localisation of m G l u R 4 and m G l u R 7 remained to be determined at the time of writing this manuscript. 75

Ch. I

J. Voogd, D. Jaarsma and E. Marani

3.4. NITRIC OXIDE: THE CEREBELLAR LOCALIZATION OF NITRIC OXIDE SYNTHASE, GUANYLATE CYCLASE AND CYCLIC GMP Nitric oxide (NO) (see Dawson et al., 1992 and Vincent and Hope, 1992 for reviews) has gained importance as an intracellular and diffusible intercellular messenger in the cerebellum, since the demonstration by Garthwaite et al. (1988, 1989) that N-methyl-Daspartate (NMDA) receptor activation caused an increase in cyclic guanosine 3',5'monophosphate (cyclic GMP) in the cerebellum by stimulating the release of a diffusible messenger with properties similar to a endothelium-derived relaxing factor which was identified as NO. They considered granule cells as the main source of NO and glial cells as the main target for the activation of soluble guanylate cyclase by NO and the production of cyclic GMP. The enzyme nitric oxide synthase (NOS), that produces NO and citrullin from arginine, occurs as several isoenzymes (Knowles et al., 1989). Type I NOS is a constitutive, calcium and calmodulin-dependent enzyme, present in neurons and, possibly, in glia. Type II NOS is calcium-independent and can be induced in macrophages and glial cells by exposure to bacterial lipopolysaccharide (Galea et al., 1992; Murphy et al., 1993). Type III NOS is the endothelial iso-enzyme. NOS-I, II and III are produced by different genes (Bredt et al., 1991; Lamas et al., 1992; Xie et al., 1992; Lowenstein et al., 1992; Lyons et al. 1992; Ogura et al., 1993). NOS displays NADPH-dependent diaphorase

Fig. 54. Ultrastructural localization of mGluR2/3 immunoreactivity in the granular layer of rat cerebellar cortex. Dense immunoreaction products accumulate in axon terminals of Golgi cells, which often make synaptic contacts (curved arrows) with possible granule cell dendrites around a mossy fiber terminal (MT) in the cerebellar glomerulus. Bar = 0.5 r Ohishi et al. (1994).

76

The cerebellum." chemoarchitecture and anatomy

Ch. I

activity and can be demonstrated in aldehyde-fixed tissue by NADPH-dependent reduction of tetrazolium salts to visible formazans (Hope et al., 1991). NOS-I has been localized with antisera to the purified enzyme (Bredt et al., 1990) and by in situ hybridization to NOS-I mRNA (Bredt et al., 1991) in basket cells and in granule cells and their axons, where NOS-I is co-localized with NADPH diaphorase (Bredt et al., 1991; Vincent and Kimura, 1992; Schmidt et al., 1992; Schilling et al., 1994). NADPH-diaphorase-positive granule cells are distributed in a symmetrical pattern of heavily stained clusters, separated by granule cells that were stained weakly, or not at all (Fig. 55) (Schilling et al., 1994). The NADPH-diaphorase-positive granule cell clusters were correlated with the Zebrin pattern in the overlying molecular layer by Hawkes and Turner (1994). A sparse axonal network and a few cells were stained in the cerebellar nuclei (Vincent and Kimura, 1992). Schmidt et al. (1992) also found weak NOS-I immunoreactivity in Bergmann glia and astrocytes where it co-localized with NADPH-diaphorase. NOS-II was expressed by astrocytes in lipopolysaccharide-stimulated cultures. These cells also double-label for NADPH-diaphorase (Galea et al., 1992). Guanylate cyclase, the enzyme responsible for the synthesis of cyclic GMP from guanosine triphosphate, was localized with immunofluorescence in Purkinje, granule stellate and Golgi cells and in oligodendrocytes, astroglia and Bergmann glial fibers of the cerebellar cortex of the rat (Zwiller et al., 1981). The localization in Purkinje and granule cells and in astrocytes was confirmed by Ariano et al. (1982), Nakane et al. (1983) and Schmidt et al. (1992). Bergmann glia and small cells in the molecular and granular layers were weakly stained. Expression of the soluble guanylyl cyclase mRNA in rat cerebellum was moderate in Purkinje, basket, stellate and Golgi cells, weak in granule cells, but could not be demonstrated in glial cells (Matsuoka et al., 1992, see also Burgunder and Cheung, 1994). Cyclic GMP was located with immunohistochemical methods in Bergmann glia (Cumming et al., 1977, 1979; Chan-Palay and Palay, 1979; Ariano et al., 1982) and in a subpopulation of stellate and basket cells (Chan-Palay and Palay, 1979). Its preferential localization in Bergmann glia and cerebellar astrocytes was stressed by Berkelmans et al. (1989) and De Vente et al. (1989, 1990), using antibodies against conjugates of cyclic GMP and activation of cyclic GMP by sodium nitroprusside in slices of rat cerebellum. They observed a patchy distribution of the reactive Bergmann glia in the molecular layer (Fig. 56). Purkinje and granular cells remained unstained. Immunoreactive varicose (mossy?) fibers and astrocytes and/or Golgi cells were observed in the granular layer. Owing to the differential localizations of NOS, guanylate cyclase and cyclic GMP, the cellular basis for the actions of cerebellar NO remains difficult to establish. Basket and stellate cells appear to be the only cell types that can be stimulated by NMDA receptors (Quinlan and Davis, 1985; Hussain et al., 1991) that contain both NOS-I, guanylate cyclase and cyclic GMP. It has been suggested that carbon monoxide (CO) is an activator of soluble guanylyl cyclase in Purkinje cells. Heme oxygenase-2, which degrades heme to biliverdin and releases carbon monoxide in the process, was shown to be co-localized with guanyl cyclase in rat Purkinje and granule cells with in situ hybridization histochemistry (Verma et al., 1993). 3.5. ADENOSINE, 5'-NUCLEOTIDASE AND ADENOSINE DESAMINASE Adenosine-like immunoreactivity was found in rat Purkinje cells, using polyclonal anti77

Ch. I

J. Voogd, D. Jaarsma and E. Marani

Fig. 55. Coronal section through the copula pyramidis (lobule VIII). In the adult rat granule cells in the lateral tip of the copula pyramidis show strongly reduced staining intensity for NADPH-diaphorase, in contrast to the medial copula, where a cluster of heavily stained granule cells can be seen. g, granular layer; m, molecular layer. Scale bar = 200/lm. Schilling et al. (1994).

sera against a conjugate of the adenosine derivative laevulinic acid (Braas et al., 1986). Staining was present in the cell soma outside the nucleus, extending in the dendrites. Weaker staining was observed in the granular layer. Adenosine is co-released with adenosine triphosphate (ATP) and certain neurotransmitters (Richardson and Brown, 1987). High affinity uptake sites for adenosine are present in all layers of the cerebellar cortex (Marangos et al., 1982; Nagy et al., 1985; Biss6rbe et al., 1985). Steady state concentrations of adenosine are maintained through the activities of only three enzymes, 5'-nucleotidase (5'-N), adenosine kinase and adenosine deaminase. Adenosine kinase and adenosine deaminase were located mainly in the soluble fractions of rat cerebellar homogenates, whereas 5'-N was present in subcellular fractions (Philips and Newsholme, 1979), mainly in the synaptosomal fraction (Marani, 1977). Adenosine deaminase-immunoreactivity in rat cerebellum was present with one out of five polyclonal sera prepared by Nagy et al. (1988). Staining was present in most Purkinje cells with a variation in intensity. Staining was observed in the Purkinje cell axons and terminals in the cerebellar and vestibular nuclei. The localization of 5'-N will be discussed below. Adenosine blocks the parallel fiber-induced simple spike discharge in Purkinje cells (Kostopoulos et al., 1975) but not the climbing fiber-mediated synaptic transmission (Kocsis et al., 1984). The effect of adenosine is presynaptic and is mediated by A1adenosine receptors that are located on parallel fibers. A 1-adenosine receptors are coupled to pertussis toxin-sensitive G proteins and inhibit adenyl cyclase. Activation of A 1-adenosine receptors decreases transmitter release from the terminals (Dolphin and Prestwich, 1985, see Fredholm and Dunwiddie, 1988, for a review). The presence of Al-adenosine receptors on parallel fibers was demonstrated autoradiographically by 78

The cerebellum." chemoarchitecture and anatomy

Ch. I

Fig. 56. cGMP-immunostaining of adult rat cerebellum. A. Section of a cerebellar slice that was incubated with cyclic GMP antiserum, in the presence of 1 mM isobutyl-methylxanthineto inhibit phosphodiesterase activity and 10 r nitroprusside and post-fixed in paraformaldehyde. B. Same areas of the same section as shown in (A) after removal of cGMP-immunostaining using the methods of Tramu et al. (1978) and reincubation of the section with glial fibrillary acidic protein-antiserum. Note presence of cGMP-immunoreactivity in Bergmann glial fibers and in thin varicose fibers (arrows in A) and in astrocytes or Golgi cells (arrow head in A). Bars = 100 r De Vente et al. (1989).

Goodman and Snyder (1982) and Goodman et al. (1983) using specific binding of [3H]cyclohexyladenosine ([3H]CHA) and Weber et al. (1990), using the antagonist [3H]DPCPX (Fig. 57). Binding was highest over the molecular layer, with lower concentrations in the granular layer. Binding was absent in the granuloprival cerebellum of 'weaver' mice (Goodman et al., 1983; Wojcik and Neff, 1982, 1983). Al-adenosine receptors were present over the entire molecular layer; no bands of high activity, corresponding to the 5'-N pattern, were observed (Fastbom et al., 1987). Adenosine is released in a Ca2+-dependent manner by K + stimulation from rat cerebellar slices (Cu6nod et al., 1989; Do et al., 1990). The stimulated release of adenosine was decreased by 60-70% in vermis and hemisphere, in slices from 3-acetylpyridine-treated rats, which may indicate that the released adenosine, at least in part, is released by climbing fibers. The 'climbing fiber-dependent' adenosine release, however, occurs with some time delay after the K + stimulus. Adenosine, therefore, has been proposed to be derived from extracellular degradation of released nucleotides by ectonucleotidases. Inhibition of 5'-nucleotidase (5'-N) by ~,fl-methylene-ADP and GMP, indeed, decreased stimulated adenosine release by 50-60%. 5'-Nucleotidase (5'-N) is an integral glycoprotein of the cellular plasma membrane in a wide range of animal cells. Its functional role is still unclear. 'Possibilities .... include recovery of purines and pyrimidines from the extracellular space, the extracellular formation of neuromodular adenosine from released nucleotidases and non-enzymatic functions related to the interaction of 5'-nucleotidase with compartments of the cytoskeleton and extracellular matrix' (Schoen et al., 1987). 5'-N catalyses the production of adenosine by the hydrolytic cleavage of 5'-nucleotide monophosphates (i.e. adenosine5'-monophosphate). The development of 5'-N in the cerebellum was studied by Schoen et al. (1987, 1988, 1990). 5'-N in the molecular layer of mouse cerebellum is distributed in positive and negative parasagittal bands (Scott, 1963). The distribution of cerebellar 5'-N has been reviewed by Marani (1986). Its zonal distribution in mice is very similar to the distribution of the m a b Q l l 3 (Zebrin)-positive dendrites of Purkinje cells in the molecular layer (Marani, 1986; Eisenman and Hawkes, 1989) (Figs 58A, 130, 131, 135) (Section 6.1.4.). The 79

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to a section of rat cerebellum. Left. Photomicrograph of a pyrorine Y-stained section of rat cerebellum showing the molecular layer (ML), the Purkinje cell perikarya (PC), the granule cell layer (GL), and some white matter ( W M ) . R i g h t . Darkfield photomicrograph of the tissue section incubated with 0.8 nM [3H]DPCPX and apposed to nuclear emulsion-coated coverslips. The silver grains are from the same area shown on the left. Note the high density of A~ adenosine receptors in the molecular layer and moderate labelling in the granule cell layer. The white matter and the Purkinje cell bodies showed background levels of labelling. Bar = 5 0 / l m . Weber et al. ( 1 9 9 0 ) .

reaction product in the histochemical procedure of Scott (1964, 1965, 1967) is uniformly distributed within the bands of high 5'-N activity in the molecular layer. In the hemisphere the staining in some of the 5'-N bands is less uniform and assumes the aspect of radially disposed striations. Hess and Hess (1986) tentatively identified these striations as the processes of the Bergmann glia. These authors found 5'-N in the molecular layer of Purkinje cell-deficient mice (pcd and nr strains) to be reduced and the residual enzyme activity to be localized in approximately the same position as the surviving Purkinje cells. This would imply that the expression of 5'-N in Bergmann glia is regulated by the adjoining Purkinje cells. Marani's electron microscopic enzyme-histochemical studies (Marani, 1981, 1982a,b, 1986), favoured a localization of the enzyme in the subsurface cisterns and the spine apparatus of the spines of the Purkinje cell dendrites and within boutons of parallel fibers (Fig. 58B-D). A non-zonal distribution of 5'-N in the somata of Purkinje cells and other large cells of the cerebellar cortex was observed when different substrates for the enzyme histochemical reaction for 5'-N were used (Scott, 1967; Marani, 1982a and b, 1986; Hess and Hess, 1986). According to Marani this represents a rest-activity of non-specific phosphatases, that disappears when the appropriate inhibitors are used. The presence of 5'-N in parallel fibers was disputed by Hess et al. (1983), who showed that 5'-N remains at significant levels in the molecular layer in agranular 'weaver' 80

The cerebellum." chemoarchitecture and anatomy

Ch. I

cerebellum. According to Kreutzberg et al. (1978) 5'-N is predominantly associated with glial membranes. Schoen et al. (1987, 1988), used monoclonal and polyclonal sera directed against rat liver 5'-N in the localization of cerebellar 5'-N in addition to the enzyme-histochemical techniques. They found the enzyme to be situated at the outer border of the plasma membranes of Bergmann glial fibers in the molecular layer, astroglial endfeet around blood vessels and glial processes surrounding Purkinje and granule cells (Fig. 59). They were unable to confirm Marani's observations of an intracellular localization of the enzyme. The study of Schoen et al. (1987) was done in rats, which do not have the longitudinal band pattern of 5'-N with their antibody directed against this enzyme. Balaban et al. (1984) observed an increase of cerebellar 5'-N in the P2 (synaptosome) fraction after climbing fiber activation with harmaline in rats (Fig. 60). Harmaline synchronizes the discharge in climbing fibers from certain parts of the inferior olive and induces a rhythmic tremor (Sj61und et al., 1977, 1980). Two different climbing fiber induced effects, therefore, may be involved in adenosine-mediated blockade of transmission in parallel fiber-Purkinje cell synapses: an increased release of nucleotides and an increase of cerebellar 5'-N. Loss of climbing fiber-induced 5'-N and/or adenosinemediated blockade of transmission in the parallel fiber-Purkinje cell synapses (see Marani, 1986) would explain the long-term increase of simple spike activity that occurs when complex spikes are suppressed by destruction or inactivation of the inferior olive in rats (Colin et al., 1980; Montarolo, 1982). Bloedel and Lou (1987), however, observed a short-term facilitation of transmission in the mossy fiber-parallel fiber-Purkinje cell pathway on stimulation of climbing fibers in the cat. This difference may be due to species-dependent differences in 5'-N mediated formation of adenosine or to a facilitation at the level of mossy fiber-granule cell synapse. If the formation of adenosine is largely dependent on the degradation of nucleotides by 5'-N, the zonal distribution of this enzyme in different species and of the climbing fibers which promote their release would be of crucial importance (see Marani (1986) and Section 6.1.4.). 3.6. INTERNEURONS OF THE CEREBELLAR CORTEX Stellate, basket and Golgi cells are inhibitory (Eccles et al., 1964a, 1966a,b,c,d, 1967). It was against this background that Uchizono (1965) (see also Uchizono, 1969 for a review) formulated and tested his hypothesis that excitatory and inhibitory axon terminals in aldehyde fixed tissue can be distinguished by the shape of their synaptic vesicles (Fig. 61). Inhibitory boutons contain flattened vesicles (F-type boutons) and excitatory boutons contain spherical vesicles (S-type boutons). Earlier Gray (1959) distinguished two types of synaptic junction, which were also supposed to represent the excitatory and inhibitory synapse (Landis and Reese, 1974). Gray type 1 junctions are characterized by a widening of the synaptic cleft that contains dense material and a distinct asymmetry caused by the presence of a dense undercoating of the postsynaptic membrane. It was considered to be excitatory. The thickening of the pre- and postsynaptic membranes in the Gray type 2 junction is symmetrical and the cleft is narrow; this type was supposed to be inhibitory. According to Uchizono (1969) there is an excellent correlation in the cerebellar cortex of the cat of S-type boutons with Gray's type 1 synaptic junctions and of F-types with a synapse of Gray's type 2. For the excitatory connections of the mossy and climbing fibers and for the parallel fiber-Purkinje cell synapse the correlation with S-type terminals and Gray 1 synaptic junctions still is valid. For the terminals of the inhibitory interneurons of the cerebellar cortex (Golgi cells: pleomorphic vesicles, synap81

Ch. I

J. Voogd, D. Jaarsma and E. Marani

Fig. 58. Light and electron micrographs of incubations for 5-nucleotidase according to Scott (1967). A. Detail of the light microscopic location of 5'-nucleotidase in uvula (IX) and pyramis (VIII). B. Electron microscopic location of 5'-nucleotidase reaction products in the subsurface cisternae of a Purkinje cell dendrite. C. Electron microscopic localization of 5'-nucleotidase in the spine apparatus of Purkinje cell dendritic spines (asterisks). D. Localization of reaction product in a parallel fiber bouton, synapsing on a Purkinje cell dendritic spine. Bars in A = 1 mm, in B,C = 0.5 ~tm, in D = 0.25 ~tm. Marani (1977).

82

The cerebellum. chemoarchitecture and anatomy

Ch.I

Fig. 59. 5'-Nucleotidase immunohistochemical staining of rat cerebellum. A. Immunofluorescence. B. PAPmethod. Enzyme activity is predominantly found within the molecular layer on Bergmann glial fibers (long arrows). Purkinje cells are surrounded by fine rims of reaction product (small arrows). Within the granular layer 5'-nucleotidase activity is diffusely scattered between granule cells (arrow heads). Vibratome sections. C. Longitudinally sectioned Bergmann glia cell processes (B) of the molecular layer of rat cerebellum. Fine DAB reaction product is located on adjacent membranes of these processes (arrows). Bars in A,B = 50 ~tm, in C = 0.5/lm. Schoen et al. (1987).

tic junction resembles Gray type 1; basket cells: ellipsoid, irregular and spherical vesicles, Gray type 2; stellate cells: flattened vesicles, Gray type 2) there is a greater variation in morphology (Palay and Chan-Palay, 1974). Cell bodies and terminals of the Golgi, basket and stellate cells can be labelled with selective uptake of [3H]GABA (H6kfelt and Ljungdahl, 1970, 1971; Schon and Iversen, 1972), immunostaining with antibodies against GAD (Saito et al., 1974; McLaughlin et al., 1974; Oertel et al., 1981b; Mugnaini and Oertel, 1985) and in situ hybridization for G A D 6 5 and G A D 6 7 (Wuenschell et al., 1986; Julien et al., 1987; Esclapez et al., 1993; Feldblum et al., 1993). They are also immunostained with antibodies against conjugates 83

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Fig. 60. Harmaline-induced changes in 5'-nucleotidase (5'-N) activity of cerebellar fractions in rats with an intact inferior olive (vehicle injection on day 1) or a 3-acetyl-pyridine (3-AP) olivectomy. Data are represented as means + S.E. for 8 animals in each group. The changes in 5'-N levels in each fraction (CH-crude homogenate, P 1, P2, P3 and $3) are shown for intact and 3-AP olivoectomized animals that served as either controls (C) or received harmaline injections (H) 45 min prior to decapitation. Harmaline evoked an increase in 5'-N activity in the CH and P2 fraction of rats with an intact olive; it evoked a decrease in the activity after 3-AP olivectomy. No significant effects appear in the P1, P3 or $3 fractions in either intact or 3-AP olivectomized animals. Balaban et al. (1984).

of GABA (Figs. 15, 40, 62 and 63) (Ottersen and Storm-Mathisen, 1984a,b; Somogyi et al., 1985; Aoki et al., 1986; Gabbott et al., 1986; Matute and Streit, 1986; Ottersen et al., 1987). 3.6.1. Stellate and basket cells Stellate cells are located in the entire, and basket cells in the deep part of the molecular layer. The dendritic arborizations of both cell types are flattened in a plane perpendicular to the long axis of the folium. Both receive synapses from parallel fibers on their dendrites. The axon of the stellate cell terminates on shafts of dendrites from Purkinje, basket, Golgi and stellate cells in the molecular layer. The immunoreactivity of stellate cells for antibodies against conjugates of taurine (Madsen et al., 1985; Magnusson et al., 1988; Ottersen etal. 1988b) is low. This is in contrast with the selective uptake by stellate and basket cells of [3H]taurine and the immunoreactivity of these cells with antibodies against CSADS, the synthesizing enzyme of taurine (Chan-Palay et al., 1982a,b) (see Section 3.1.2.). The localization of CSADS in basket and stellate cells has, however, been disputed, since Almerghini et al. (1991) found CSADS-immunoreactivity to be exclusively localized in glial cells and not in neurons of the cerebellar cortex. Basket axons have been studied extensively (Palay and Chan-Palay, 1974). They extend in a direction across the axis of the folium (Fig. 13) and terminate with ascending 84

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Fig. 61. Size and shape analysis of synaptic vesicles in S-type and F-type synapse. Diameters of both major and minor axis in each synaptic vesicle in both types of synapses were measured. Ordinate shows the length of the major axis, while the abscissa that of the minor axis of vesicles in each type of synapse. Diameters of vesicles in S-type synapse (white) are distributed around the 45-degree line between ordinate and abscissa, while those in F-type synapse (black) of both white and black circles indicates the relative frequency of occurrence. Elongation index ratio of average length of major versus minor axis of vesicles in S-type synapse was about 1.2, while that of F-type synapse was about 1.7. Uchizono (1965).

branches on the primary dendrites of Purkinje cells and constitute the baskets surrounding the cell body that end in the pinceau around the initial part of the axon of the Purkinje cells. GABA-like immunoreactivity was present in boutons of stellate and basket cell axons on Purkinje cell dendritic shafts, in basket cell terminals on dendrites of stellate cells and on Purkinje cell somata (but not in all of them) and in some of the axons of the pinceau (Gabbott et al., 1986). No specific neurochemical properties seem to distinguish the basket cells from the stellate cells. According to Somogyi et al., (1986) GABA-like immunoreactivity is weaker in stellate cells than in basket and Golgi cells. Basket and stellate cells are immunoreactive for antibodies against parvalbumin, like the Purkinje cells (Fig. 31B). No reactivity for these antibodies or m R N A probes for parvalbumin was mentioned for the Golgi cells (Celio and Heizmann, 1981; Heizmann, 1984; Schneeberger et al., 1985; Endo et al., 1985; Braun et al., 1986; Kadowaki et al., 1993; Kosaka et al., 1993). Calretinin immunoreactivity is present in a subpopulation of stellate and basket cells in the cerebellum of the chicken, where it is co-localized with parvalbumin in some of the cells (Rogers, 1989). Immunoreactivity with antibodies against PKC ~, fl, g, e and possibly ~"is present in stellate and basket cells (Fig. 29, Table 1). The localization in basket and stellate cells of nitric oxide synthase, guanylyl cyclase and cyclic GMP has been reviewed in Section 3.4.

3.6.2. Golgi cells and Lugaro cells Golgi cells are located in the granular layer, and have been roughly subdivided into large Golgi cells, which are located in the superficial part of the granular layer and small Golgi cells located more deeply. The dendrites of both types extend into the molecular layer, where they are not confined to a single plane. The axons branch repeatedly to form a 85

Ch. I

.

9.t ; . ~ ~

.....

J. Voogd, D. Jaarsma and E. Marani

.

*~

";Y~'

"~

~j~!~;i;i:..:k!~,~.~ ~:, :::~ . .

'

"

~

.... ~ , ~ o

.: ~

~..~

~ ......

.............. ~ - . ~ . ~. . . . . . . . . . . . . ~ .....................

~. ...,

...

Fig. 62. Cerebellar cortex of cat reacted by the unlabelled antibody enzyme method. A and B. Serial semithin (1/lm) sections reacted under postembedding conditions with an anti-GABA serum (A) or with the same serum after solid phase absorption (B). Basket cells (BC) and Golgi cells (GC) are strongly immunoreactive, Purkinje cells (PC) and stellate cells (SC) reacted less strongly. GABA-immunoreactive terminals are present in all layers, but the terminals of basket cells around the Purkinje perikarya and in the pinceau (p) and the Golgi cell terminals in the glomeruli (gl) are especially strongly reacting. C-E. Preembedding demonstration of GABA and GAD in vibratome sections. The distribution of amino acid and its synthesizing enzyme are very similar, but perikarya of basket, stellate, and Golgi cells, and basket cell axons (ba) stain stronger for the amino acid than for G A D in animals not treated with colchicine. F and G. Electron micrographs of glomeruli demonstrating GABA (F) and GAD (G) in the terminals (asterisks) of Golgi cells. The dendritic digits of granule cells (diamonds) receive synapses (arrows) from the immunoreactive terminals as well as from the mossy fiber terminals (mft). Bar in A-E - 50/~m, in F and G = 0.5 r Somogyi et al. (1985).

86

The cerebellum." chemoarchitecture and anatomy

Ch. I

dense plexus in the granular layer. The terminals participate in the formation of the glomeruli where they make synaptic contact with the granule cell dendrites. Golgi cells in the upper molecular layer in rat and cat are selectively recognized by a monoclonal antibody (rat-303, Hockfield, 1987) (Figs. 66 and 69B). Uptake studies have shown that [3H]GABA and [3H]glycine uptake result in similar patterns of axonal labelling in the granular layer, in circular deposits resembling the periphery of the glomeruli, whereas no [3H]glycine labelling was found over the pericellular baskets of the Purkinje cells (Wilkin et al., 1981a). Concordantly it was shown that a large proportion of the Golgi cells, in addition to GABA, was also immunoreactive for antibodies against conjugates of glycine (Ottersen et al., 1987, 1988a; Campistron et al., 1986a; Takayama, 1994). In a high percentage of these glycine containing Golgi cells (40%) glycine-like and GABA-like immunoreactivity co-exist (Ottersen and StormMathisen, 1987). GABA and glycine-like immunoreactivity co-exist in most Golgi cell

~:.

.. ~!

Fig. 63. Photomicrographs of semithin (0.5 r tissue and test sections incubated with GABA antiserum 26 diluted 1:100 (A) or glycine antiserum 31 diluted 1:60 (B). Three of the four Golgi neurons that are glycine immunoreactive (thick arrows) are also stained with the GABA antiserum in the adjacent section; the fourth glycine-positive neuron (crossed arrow) is virtually immunonegative for GABA. Most if not all glomeruli (arrowheads) show GABA-like-immunoreactive as well as glycine-like-immunoreactive positive Golgi cell terminals. The molecular layer contains no glycine-like immunoreactive positive structures except for a few fibrous processes (small arrows in B). The terminals of the basket and stellate cells and their respective cell bodies (double arrowhead) are glycine immunonegative, but GABA immunopositive. Asterisks, Purkinje cell bodies. Other abbreviations: MO and GC, molecular and granule cell layers, WM, white matter. Bar = 50 r Ottersen et al. (1988a).

87

Ch. I

J. Voogd, D. Jaarsma and E. Marani

Fig. 64. Electron micrograph showing GABA-LI in a glomerulus in the granular layer of the cerebellar cortex (rat). After incubation in rabbit primary antiserum, the section was treated with sheep anti-rabbit immunoglobulins bound to colloidal gold particles (16 nm). Axon terminals of the inhibitory Golgi cell (GO) show a high density of gold particles, whereas the densities of such particles over the mossy fibre boutons (MF) and granule cell dendritic digits (GC) are close to back-ground level. Scale bar = 0.5 ~tm. Ottersen and StormMathisen (1987).

terminals located at the periphery of the glomeruli (Somogyi et al., 1986; Ottersen et al., 1987) (Fig. 63) (Ottersen et al., 1988a) (compare Figs 41 and 65). A similar localization of GABA-like and glycine-like immunoreactivity was found in the rat and the baboon (Ottersen et al., 1987) (Fig. 65). Some displaced Golgi cells were present in the molecular layer of the baboon and many fibers running in the supraganglionic plexus in the direction of the long axis of the folium displayed glycine-like immunoreactivity. In summary the majority of the Golgi cells may use glycine in addition to GABA. Golgi cells in the rat stain strongly for acetylcholinesterase (Brown and Palay, 1972; 88

The cerebellum." chemoarchitecture and anatomy

Ch. I

Altman and Das, 1970). They share this property with a group of displaced Golgi cells located in the lower molecular layer in the rabbit (Ramon y Cajal, 1911; Spa~;ek, 1973). A subpopulation of Golgi cells in the cat and man, but not in rat or rabbit, is immunoreactive for choline acetyltransferase (see Section 3.10.1., Fig. 86). Certain Golgi cells in rats were found to be immunostained with antibodies against conjugates of somatostatin (Johansson et al., 1984; Vincent et al., 1985; Villar et al., 1989) (Fig. 20) or enkephalin (Schulman et al., 1981; Ibuki et al., 1988). Calcium binding proteins have not been found in Golgi cells, with the exception of a single antibody against calbindin-D28k, that stains Purkinje cells and Golgi cells in rat and human cerebellum (Garcia-Segura et al., 1984; Fournet et al., 1986), and the presence of calretinin in some Golgi cells (Arai et al., 1991; Floris et al., 1994). Of the PKC subtypes only PKC e' has been localized in Golgi cells (Wetsel et al., 1992). None of the immunoreactive subpopulations of Golgi cells seem to correspond exclusively to one of the anatomical subtypes distinguished by Palay and Chan-Palay (1974). Similarly, Golgi cell heterogeneity due to differential expression of metabotropic glutamate receptors, does not correspond to any anatomical subdivision (see Section 3.3.2.). It remains to be elucidated to what extent the different immunocytochemical markers overlap. Lugaro cells are fusiform cells located below the Purkinje cell layer, with dendrites arising from opposite poles of the cell and somata extending for long distances beneath the Purkinje cell layer. Lugaro cells are chemically distinct from Golgi cells in that they are selectively recognized by two monoclonal antibodies (cat 301 and 304, Sahin and Hockfield, 1990) (Fig. 67). Like Golgi cells they are immunoreactive for antibodies against GABA (Aoki et al., 1986). Ottersen et al. (1988a) included the Lugaro cells with the Golgi cells and found GABA-like and glycine-like immunoreactivity to be colocalized in Lugaro cells. Lugaro cells are assumed to primarily innervate the granule cells (Palay and Chan-Palay, 1974), although according to Fox (1959) their axon may enter the molecular layer. 3.6.3. Unipolar brush cells

Unipolar brush cells often have been interpreted as small Golgi cells. They are, however, non-GABAergic and non-glycinergic (Aoki et al., 1986; Mugnaini et al., 1994). Since they give rise to mossy fiber rosette-like terminals in the granular layer they are likely to be glutamatergic (Berthie and Axelrad, 1994; Rossi et al., 1995) (Section 3.2.1.). Unipolar brush cells can be distinguished from other granular layer neurons by a number of immunocytochemical markers (reviewed by Mugnaini and Floris, 1994). Following their original characterization as pale cells by Altman and Bayer (1977), they were first recognized by Hockfield (1987) using a monoclonal antibody against spinal cord gray matter, Rat-302 (Fig. 69). A study of Harris et al. (1993) showed that Rat-302 is directed against high molecular weight neurofilament protein, and that unipolar brush cells are strongly immunostained by several different antibodies against high molecular weight neurofilament protein. Unipolar brush cells are essentially unstained and moderately stained with antibodies against middle and low molecular neurofilament protein, respectively. Cozzi et al. (1989) and Munoz (1990) identified unipolar brush cells in the rat and human cerebellum, respectively, on the basis of their immunoreactivity to antisera against proteins of the secretogranin (or chromogranin) family. Unipolar brush have a relatively high density of large dense core vesicles, which in conjunction with the 89

Ch. I

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J. Voogd, D. Jaarsma and E. Marani

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,,
t

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Fig. 65. Details from sagittal vibratome section of baboon cerebellum (vermis) treated with the glycine antiserum diluted 1:250. A. One of the immunopositive Golgi cells shows a dendrite (small arrowheads) extending into the molecular layer, and a thinner process (axon?) which seems to engage in a glomerulus-like structure (large arrowhead). Note the presence of thick and thin immunoreactive fibers (thin arrows) in the deep part of the molecular layer, which also contains scattered immunoreactive neurons (thick arrow). B. Radial fibrous processes (small arrows) interpreted as Bergmann glia, show weak glycine-like immunoreactivity. Large arrow points at an immunostained Golgi cell. Asterisks indicates immunonegative Purkinje cell. C. Immunopositive Golgi-like neurons (arrows) occur in or just below the layer of the unstained Purkinje cells (asterisks). Large arrowhead indicates a stained Golgi cell slightly outside the plane of focus. Inset: Arrowhead, fusiform cell with horizontal dendrites situated directly beneath the Purkinje cells (asterisk). G, granular layer; M, molecular layer. Bar = 100 ~tm. Ottersen et al. (1987).

presence of chromogranins might indicate that they are neurosecretory of some kind of peptide (see discussion Mugnaini et al., 1994). Particularly dense immunostaining in unipolar brush cells was obtained with antibodies against calretinin (R6sibois and 90

The cerebellum." chemoarchitecture and anatomy

Ch. I .~i/./~.~...:~.!;:.~:,~.i:ii~;..:.~i:~:;~i~il;~.:ili~~:~:.i.~i~.:i:.:i.i.~:..~..i:,i:Zi:i~i,iii~' i;~.~;;.i:i~;i:i;~.~i,:;i~i~,i:i~i:. ..:"": ii~!!.! !~ ;i:~../.iii:~iI i~:i!i~:":::':;~"~"!

Fig. 66. A. Monoclonal antibody Rat-303 recognizes neurons in the granule cell layer (g), but not in the Purkinje cell (p) or molecular (m) layers of the rat cerebellar cortex. Scale bar = 100/~m. B. The morphology of Rat-303-positive neurons matches that described for Golgi II cells: a large ceil body emitting relatively stout dendrites from many points over the cell circumference (I3). Scale bar = 10 ~m. Hockfield (1987). R o g e r s , 1992; B r a a k a n d B r a a k , 1993; Floris et al., 1994). C a l r e t i n i n - i m m u n o r e a c t i v i t y in u n i p o l a r b r u s h cells was u s e d for a d e t a i l e d c h a r a c t e r i z a t i o n o f this cell type, a n d to

Fig. 67. Lugaro cells are molecularly distinct from Purkinje cells. In the pairs illustrated in A and B, the same fields were photographed under fluorescent optics separately for FITC (anti-calbinding, A) and Texas Red (Cat-301 in B). The blood vessel (triangle) passing through the field can be used to align the photographs. The cell type-specific antibody Cat-301 recognizes Lugaro, but not Purkinje cells in cat cerebellum, while anticalbindin recognizes Purkinje, but not Lugaro cells. Scale bar = 50/lrn. Sahin and Hockfield (1990). 91

Ch. I

J. Voogd, D. Jaarsma and E. Marani

Fig. 68. Immuno-electron micrograph of an unipolar brush cell stained with antiserum to calretinin. The micrograph was obtained near the surface of the immunoreacted slice, as indicated by open areas in the tissue and over the cell nucleus (stars). The unipolar cell body has an irregular contour. The nucleus (N) shows a deep indentation (T) and the cytoplasm contains an array of ringlet subunits (R). The short dendrite forms an extensive synapse (arrows) with a calretinin-negative mossy fiber ending (mf). At the aspect opposite the synapse the dendrite contains numerous mitochondria. Granule cells (CG) are immunonegative. Bar = 1/~m. Floris et al. (1994).

92

The cerebellum." chemoarchitecture and anatomy

Ch. I

study its distribution in the cerebella of different species (Fig. 68) (Floris et al., 1994; Dino and Mugnaini, in preparation). 3.7. LOCALIZATION OF GABA RECEPTORS A N D GLYCINE RECEPTORS 3.7.1.

GABA A

receptors

GABA receptors can be divided in two main classes, GABAA and GABA~ (Bowery et al., 1980; Barnard et al., 1987; Sieghart, 1989; Sivilotti and Nistri, 1993; Mody et al., 1994). G A B A A receptors are ionotropic receptors that gate chloride channels. They are inhibited by bicuculline and picrotoxin and activated by muscimol. G A B A A receptors are the target of for a variety of drugs including benzodiazepine tranquilizers, and barbiturates. Benzodiazepines bind to a modulatory site that facilitates G A B A A receptor function. Different types of G A B A A receptors can be pharmacologically distinguished based on their differential sensitivity to benzodiazepine ligands and barbiturates (Doble and Martin, 1992). Originally the distribution of G A B A A receptors was studied autoradiographically with [ 3 H ] G A B A in the presence of the baclofen and the absence CaC12 or with [ 3 H ] m u s cimol. G A B A A receptors in rat and mouse cerebellum were found to be highly concentrated in the granular layer and relatively low in the Purkinje cell and molecular layers

iii!i!i......

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k:~iiii~

~.:~:~,

!',i~;:,i~i?i;i;i?:'iii~

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~ ; : , i~: i:~i ~~' !,~' i: ~!i i'~'~"~i:.~: ;:

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". . . . . . . . .

9 ............========...=....... ==== ~ . - . . .

:...:". ' ::~::~~ '. i:

~

i

Fig. 69. A. Monoclonal antibody Rat-302 recognizes a subset of neurons, later identified as unipolar brush cells, restricted to the granular layer of the flocculus and the vermis of rat cerebellum (arrows), whereas in other areas of the cerebellum no positive cells are found. B. In contrast, antibody Rat-303 recognizes Golgi-II cells in the granular layer (g) in the entire cerebellum. C. Rat-302 also recognizes Purkinje cells outside the caudal vermis and the flocculus. D. Rat-302 positive cells in the vermis. E and F. Unipolar brush cells recognized by Rat-302 have a round cell body and short dendrites ending in a spray of appendages (arrows). g, granular layer; m, molecular layer; p, Purkinje cell layer. Scale bars: 500 p m in A and B, 50 p m in C and D, 10 p m in E and F. Hockfield (1987).

93

Ch. I

J. Voogd, D. Jaarsma and E. Marani

(Fig. 70) (Palacios et al., 1980, 1981a; Kingsbury et al., 1980; Bowery et al., 1987; Rotter et al., 1988; Wilkin et al., 1981b). Binding to benzodiazepine receptors of [3H]flunitrazepam (Young and Kuhar, 1979, 1980; Vaccarino et al., 1985) or [3H]Ro15-1788 (Schoch et al., 1985), that would be expected to follow the pattern of [3H]muscimol binding to GABAA receptors, with the highest values in the granular layer, actually is reversed, with higher binding over the molecular layer. The inverse partial benzodiazepine agonist [3H]Ro 15-4513, however, exhibit higher binding to the granular layer than to the molecular layer (Sieghart et al., 1987). [3H]Ro15-4513 binding sites in the cerebellar granular layer differ from [3H]Ro15-4513 binding sites in the molecular layer and other brain regions in that they are insensitive to diazepam and other benzodiazepine agonists (see below). Yet another type of GABAA receptor is labelled with the antagonists [3H]bicuculline methochloride and [3H]SR95531, which preferentially bind to the low-affinity GABAA sites, and which bind at low density to the cerebellum (Bristow and Martin, 1988). Autoradiographic studies with mutant mice have revealed different changes in [3H]flunitrazepam and [3H]muscimol binding: [3H]flunitrazepam binding in the molecular layer in 'Purkinje cell degeneration' (pcd) mutant mice was decreased at 45 days, after the degeneration of the Purkinje cells. A further decrease was observed at 300 days, concomitant with the loss of the granule cells. Benzodiazepine receptors, therefore, may be located on Purkinje cells, and on parallel fibers in the molecular layer (Vaccarino et al., 1985; Kahle et al., 1990). An increase of [3H]flunitrazepam binding was found in 'weaver' cerebellum, with a loss of the granule cells (Chang et al., 1980; Kahle et al., 1990). Fry et al. (1985) found [3H]flunitrazepam binding to homogenates and tissue sections of the cerebellum of 'lurcher' mutant mice, with loss of their Purkinje cells and most granule cells, to be unchanged, whereas [3H]muscimol binding was reduced. Rotter et al. (1988)concluded that the decrease in [3H]muscimol binding in 'weaver', 'staggerer' and 45 days 'pcd' mutant mice was associated with a loss of granule cells and granule cell-Purkinje cell contacts. No [3H]muscimol labelling was observed over the deep Purkinje cell clusters in 'reeler' mutants, suggesting that these sites are absent on Purkinje cells. The electron microscopical studies of Chan-Palay (1978) and Chan-Palay and Palay (1978) in the rat indicate that [3H]muscimol binding to GABAA receptors is found on the plasma membrane of Purkinje cell somata, primary dendrites and initial axonal segment, and is also present on basket cells, their axons and the pinceau formation, on stellate cells and on dendrites of Golgi and granule cells. GABA A receptor subunits

To date five subunit classes of GABAA receptors, and several isoforms of each class have been cloned: ~1-~6, fll-fl3, 7'1-7'3, ~, and el and e2 (reviewed in Wisden and Seeburg, 1992; DeLorey and Olsen, 1992; Doble and Martin, 1992). GABAA receptors are constructed as hetero-oligomeric (presumably pentameric) assemblies of subunits. The ~-subunits in particular determine the different affinities of benzodiazepine ligands (Doble and Martin, 1992; Wisden and Seeburg, 1992). The y subunits are required for benzodiazepine-sensitivity, which indicates that the benzodiazepine binding site probably resides at the interface between the ~- and y-subunits. The fl-subunits are an essential structural component of the GABAA receptors, since without fl subunits, recombinant GABAA receptors are poorly expressed. The distribution of the distinct subunit mRNAs throughout the rodent cerebellum has been investigated in many studies and has been reviewed by Laurie et al., (1992) and Persohn et al. (1992) (Table 3). Most cerebellar neurons, including basket/stellate cells, Purkinje cells, and neurons of the deep nuclei 94

The cerebellum." chemoarchitecture and anatomy

Ch. I

Fig. 70. Bright-field (A) and dark-field (B) photomicrographs of a cerebellar folium of rat cerebellum labelled with [3H]muscimol to demonstrate the GABAA receptors. Note the clustering of grains over the area corresponding to the glomeruli of the granule cell layer and the low level of autoradiographic grains over the molecular layer (m) and the Purkinje cell layer (arrows). W, white matter. Bar = 100 r Palacios et al. (1981a).

express ~1, f12 and the 72 subunit mRNAs (Fig. 71, Table 3). Many types of subunits are produced by granule cells, which express c~l, ~z6,f12, f13, ?'2, fi mRNA and possibly also ill, cz4 and 7"3 mRNA (Fig. 71). No data are available on the subunit mRNAs expressed by Golgi cells. Bergmann glial cells express distinct levels of ~z2, 7'1, and possibly fll m R N A (Fig. 72, Table 3). The distribution of G A B A A receptor subunits has also been studied immunocytochemically with subunit specific antibodies. The monoclonal antibodies bd-24, which is selective for the ~zl subunit (but does not immunoreact with the rat antigen) and bd-17, which reacts with f12 and f13, preferentially stain the granular layer in rat, cat and monkey (Fig. 73) (Richards et al., 1987; Somogyi et al., 1989; Baude et al., 1992), and conform to the distribution of [3H]muscimol binding. A similar pattern was obtained with the monoclonal antibody 62-3G1 of De Bias et al. (1988), which also react with f12 and f13, and with several antibodies specific for the cz1 subunit (Meinecke et al., 1989; Gutidrrez et al., 1994 and references therein). Immunostaining in the granular layer prevails in the glomeruli, but granule cell membranes are also lightly stained (Fig. 73). Golgi cells were found to be unstained. Immunostaining in the molecular layer is compatible with a localization on Purkinje cell dendrites and stellate and basket cell somata. Purkinje cell somata and the basket cell-pinceau formations are lightly stained or unstained. In the deep nuclei, the majority of the cell bodies and also the dendrites were outlined by immunoreactivity. Electron microscopy by Somogyi et al. (1989) 95

Ch. I

J. Voogd, D. Jaarsma and E. Marani

TABLE

3.

Distribution of GABAA receptor subunit mRNAs in rat cerebellum

Subunit

Cell type PC

GrC

al

++

a2

.

a3

-

o~4

.

.

~5

.

.

a6

-

fll

.

.

++ -

yl ?'2 y3

Based

-

not detected; stellate

on data

+

-

.

.

.

or +

+

-

-

. . -

.

DCN

or +

.

-

.

or +

++

+

-

+

+

-

-

+

-

-

-

+

-

++

+

+

-

++

-

-

-

-

-

-

++

-

_

_

or +

or +

+, positive;

++, strongly

cells; Bg, Bergmann

glia; DCN,

from

Bg

.

++

f13

Symbols:

. -

f12

cells; StC,

++ .

BC/StC

Laurie

et al., 1992; Persohn

positive; deep

PC, Purkinje

cerebellar

cells; GrC,

granule

cells; BC, basket

nuclei.

et al., 1992.

showed that both bd-24 (al) and bd-17 (,82/3) immunoreactivity occurs at the synapses established by Golgi cells with granule cell dendrites. Immunoreactivity, however, is also present at non-synaptic sites throughout the surface of the granule cell including the somatic membrane which does not receive synapses, but is absent from the postsynaptic membrane specialisations of mossy fiber-granule cell synapses. No immunoreactivity is present in the Bergmann glia and granular layer astroglia. In Purkinje cells, the synaptic junctions formed by basket cells are often immunopositive. Immunoreactivity was also found along most of the dendritic surface including the dendritic spines. No bd-24 and bd-17 immunoreactivity could be observed at the parallel fibers synapses (Somogyi et al., 1989). A different subcellular localization than that of the a l and fl2/3 subunits was observed for the a6 subunit: Baude et al. (1992) showed that a6 immunoreactivity, which was confined to the glomeruli in the granular layer, was detectable only at the postsynaptic membranes facing Golgi cell boutons, and did not occur on extrasynaptic membranes. g-subunit immunoreactivity, which was also specifically localized in the granular layer, was found in both the glomeruli and on granule cell bodies (Benke et al., 1991a). Immunocytochemical studies with antibodies specific for the y2 subunit (Benke et al., 1991 b) or for each of the two splice variants, ~ ' 2 L ( o n g ) and ) " 2 S ( h o r t ) (Guti6rrez et al., 1994), showed that 7'2 immunostaining is most abundant over the molecular layer, moderately dense over the deep nuclei, and low over the granular layer. This distribution is similar to the distribution of [3H]flunitrazepam binding sites, which is consistent with the idea that the 7' subunit is required for the binding of [3H]flunitrazepam and other benzodiazepine agonists, y2L and y2s immunoreactivities seem to be similarly distributed, but their exact cellular localisation remains to be described. The subunit compositions of functional GABAA receptors in the different cerebellar cells have been discussed by Laurie et al. (1992) and Pershon et al. (1992). Purkinje cells are likely to contain alf127'2 GABAA receptor/channel complexes, and possibly also 96

The cerebellum. chemoarchitecture and anatomy

Ch. I

~6

,13 ilili-l~1ill :lllll!llll ::l ll l li:lii:~l:l"

i........

~li

Fig. 71. Bright-field photomicrographs showing cellular distribution of mRNAs of ~1, ~6, f12, f13, y2 and GABAA receptor subunits in the cerebellum of the rat. Arrows indicate examples of labelled stellate/basket cells; arrowheads delineate Purkinje cells. Gr, granule cells; Mol, molecular layer; P, Purkinje cells. Scale bar = 50 r Laurie et al. (1992).

~lf13y2 or otlfl2fl3y2 receptors. These receptors would have a B E 1 subtype of benzodiazepine site, which is consistent with the observations that BE1, but not B E 2 benzodiazepine receptors, are found at high levels in the cerebellum (Doble and Martin, 1992). ~ l f 1 2 y 2 G A B A A Receptors are also likely to be present in basket/stellate cells and neurons of the deep nuclei. Granule cells producing at least six different subunits may have multiple subtypes of G A B A A receptors. The high level of [ 3 H ] m u s c i m o l and [ 3 H ] R o 15-4513 binding in combination with a low level of [3H]flunitrazepam binding to the granular layer can be explained by the presence of o~6(fl2and/orfl3)y2 receptors, which exhibit benzodiazepine agonist-insensitive Ro 15-4513 binding as well as high-affinity [3H]muscimol binding. It has been recently demonstrated that an alcohol-non-tolerant rat line, which is highly susceptible to impairment of postural reflexes by benzodiazepine agonists, has a point 97

Ch. I

J. Voogd, D. Jaarsma and E. Marani

i~}ii!::i~!i'iiiiili~ii]i!ii!~ili!iiiiiii::ili~ii::i~tliii:t ) 'ill

i

~t:, ....................... i~ii:~'~!:::.i";i::., " ,:~ ......

~9

,

.........

9

..............

!

J......................................................... :.~ :'':

~:':::::::::..t........:.. ~

..... ~:::

i,~,, .....

................. 9

....

,., ,:::,,~:, ....... ~:,,,,~

~

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~:,s

':;:::

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" " " ~Ji:"!~ . ~

~

~

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Fig. 72. ~2 (A) and 7'1 (B and C) GABAA subunit mRNAs are localized in Bergmann glial cells in rat cerebellum. A and B are low-power dark-field, C is high-power bright field of the image in (B)./3, putative Bergmann glia; Gr, granule cells; Mol, molecular layer; P, Purkinje cells. Arrowheads in A and B indicate 'halo' of silver grains along the boundary of the granule cell/molecular layers. In a high-power bright-field view of the 7'1 probe autoradiographic signal (C), Purkinje cells (arrow heads) and granule cells appear to be unlabelled, whereas other small cells (arrows) in the Purkinje layer have clusters of silver grains over them. There is also a density of grains higher than background over the molecular layer in areas having no cell bodies. Scale bars in A and B = 100 ~m, in C = 35/lm. Laurie et al. (1992).

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Fig. 73. Low-power dark-field (A) and microscopic brigh-field (B, Normarski optics) images of the distribution of immunoreactivity in the rat cerebellum using mAb bd-17 against the fl2/3 GABA A receptor subunits. Immunostaining is very intense over the granular layer (PMgr), moderately dense over the molecular layer (PMmo) and the deep nuclei and absent in the white matter (my). In B, note the intense staining of the glomeruli in the granular layer, and virtually absence of staining in Purkinje cells (Pc). Bars 1 mm in A, 50/~m in B. Richards et al. (1987).

99

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mutation in the ~6 subunit, which dramatically increases the sensitivity of ~6fl27'2 receptors for benzodiazepine agonists (Korpi et al., 1993). Since ~6 subunits are exclusively expressed by cerebellar granule cells these data indicate that the increased susceptibility to benzodiazepine impairment of postural relexes is mediated at the level of the granule cells, and imply that ~6 subunit containing G A B A a receptors are important in the cerebellar circuits controlling movements (Korpi et al., 1993). Immunoprecipitation indicate that ~6 mostly colocalize with ~1, f12 or/33, and 7'2 (Khan et al., 1994). It has also been proposed that ~6 assemble into GABAA receptor complexes without ~1, and into receptors without 7'2, but with the g subunit (see Laurie et al., 1992). Granule cells may also make G A B A A receptors without the e6 subunit. Mfiller et al. (1994) characterized the properties of GABAA receptors in Bergmann glial cells in developing rat cerebellum. Significant benzodiazepine-insensitive GABAA receptor conductancies were detected in Bergmann glial cells between post-natal days 7 and 10, but were small or undetectable in the adult. It was also observed that Bergmann glial cells in addition to c~l-immunoreactivity, transiently express g-subunit immunoreactivity, indicating that the transient GABAA responses are mediated by a GABAA receptor composed of ~1 and ~ subunits. In view of the in situ hybridisation data also fll and yl may contribute to the Bergmann glial GABAA receptor (Laurie et al., 1992; Pershon et al., 1992).

3.7.2. GABAB receptors GABAB receptors are insensitive to the GABAA-antagonists bicuculline or picrotoxin, and are selectively activated by baclofen. GABAB receptors are believed to be G-protein coupled receptors similar to the metabotropic glutamate receptors. Their molecular structure remains, however, to be elucidated (Bowery, 1993). GABAB receptors are mostly coupled to adenylate cyclase, and exert their inhibitory action through the activation of potassium channels or inhibition of CaZ+-channels (Bowery, 1993). Ca 2+sensitive binding of [3H]GABA, with suppression of GABA A binding by isoguvacine, or binding of [3H]baclofen were used to study the distribution of GABAB receptors. GABAB sites are present at high density in the molecular layer (Wilkin et al., 1981b; Gehlert et al., 1985; Bowery et al., 1985, 1987), where they are distributed in a pattern of alternating, parasagittal zones of high and low [3H]GABA binding (Albin and Gilman, 1988; Turgeon and Albin, 1993). GABAB receptor binding in the granular layer is very low, but above background (Turgeon and Albin, 1993). A low amount of GABAB receptor binding was found over the cerebellar nuclei in adult rats. The cerebellar nuclei, however, transiently express a very high density of GABAB receptors during development (Turgeon and Albin, 1993). The cellular localization of GABAB receptor binding in the molecular layer is controversial (reviewed in Turgeon and Albin, 1993). Wojcik and Neff (1984) reported that GABAB induced inhibition of adenylate cyclase was reduced in 'weaver' mutant mice with a loss of their granule cells. It was not affected by Purkinje cell loss in the appropriate mutants or by climbing fiber deafferentation by 3-acetylpyridine in rats. Kato and Fukuda (1985) found [3H]baclofen binding to GABAB receptors to be decreased in homogenates of rat cerebellum, after destruction of the inferior olive with 3-acetylpyridine. They concluded that GABAB receptors were located on climbing fibers and that the residual binding could be associated with granule cells. They explained the apparent discrepancy with the results of Wojcik and Neff (1984) by assuming that the high affinity binding sites, which were mainly affected in their experiments, were not 100

The cerebellum." chemoarchitecture and anatomy

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coupled to adenylate cyclase and, therefore, would have escaped to be noticed in the experiments of these authors. Bowery et al. (1987), who analyzed GABAB binding in several strains of mutant mice, however, advocated that the majority of GABAB receptors had a postsynaptic localization on Purkinje cell dendrites. Turgeon and Albin (1993), who analyzed GABAB receptor binding in rats with methyl azoxymethanol lesions of granule cells, or with 3-acetylpiridine lesions, and in stumbler mutant mice lacking Purkinje cell dendrites, also concluded that GABAB sites are primarily located on Purkinje cell dendrites. Finally, a recent immunoelectron microscopic study with monoclonal antibodies to L-baclofen indicate that GABAB receptors are found on both Purkinje cell dendrites and parallel fibers (Martinelli et al., 1992).

3.7.3. Glycine receptors Glycine-mediated inhibition of neuronal activity results from activation of the glycine receptor, a ligand-gated chloride channel (Betz, 1991). The distribution of this receptor has been autoradiographically studied with the antagonist [3H]strychnine. These studies indicate that the glycine receptor is almost absent from the cerebellum (Zarbin et al., 1981). The cerebellum, however, displays a significant number of [3H]glycine binding sites mainly distributed over the granular layer (Bristow et al., 1986), but these sites are assumed to represent the glycine coagonist sites of NMDA receptors (see Section 3.3.1 .). The glycine receptor has been demonstrated to be a pentameric protein composed of ligand binding ~ and structural fl subunits (Langosch et al., 1990). Variants of the ligand-binding ~ subunit (~1, ~2, ~3) have been cloned, that modify the pharmacological and the physiological properties of the glycine receptors. In situ hybridisation has revealed a hybridisation signal of the probe for the/3 subunit in all layers of the cerebellar cortex and the cerebellar nuclei, with a particular strong signal in the Purkinje cell and granular layers (Malosio et al., 1991). The ~l-subunit mRNA is expressed at low level in the cerebellar nuclei, and by 'rare single cells' in the granular layer, whereas ~3 mRNA appears to be selectively expressed by cerebellar granule cells. The ~2-subunit mRNA is not produced in the adult cerebellar cortex (Malosio et al., 1991). A monoclonal antibody selective for the N-terminal sequence of the ~ subunits (mAb2b), immunostained sparse puncta on cell somata in both the granular and molecular layer (Kirsch and Betz, 1993). Another monoclonal antibody, mAb4a, which bind to ~ and fl subunits, in addition to sparse puncta in the granular and molecular layer also produced diffuse labelling over Purkinje cell perikarya and occasional cell bodies in the granular layer (Kirsch and Betz, 1993). The punctate labelling is believed to reflect subunit staining at post-synaptic membrane specialisations (Kirsch and Betz, 1993). The diffuse labelling, instead, probably represents the fl subunit. Gephyrin is a 93 kDa peripheral membrane protein that co-purifies with glycine receptors. It has been proposed to anchor the glycine receptor to sub-synaptic tubulin (Betz, 1991). In situ hybridisation (Kirsch et al., 1993) and immunocytochemistry (Araki et al., 1988; Kirsch and Betz, 1993) show that gephyrin is widely distributed throughout the brain, including the cerebellum. Gephyrin mRNA is expressed by Purkinje cells and granule cells. Punctae of gephyrin immunostaining were concentrated over the molecular layer, and on cell bodies in the granular layer, but were virtually absent on Purkinje cell somata. The functional implications of the presence of glycine receptor subunits and gephyrin in the cerebellum are poorly understood. Granule cells express 0~3, the fl subunit and gephyrin and, therefore, have all the ingredients to produce functional glycine receptors. 101

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These receptors could be involved in the inhibitory actions of Golgi cells, in view of the fact that a subpopulation of Golgi cells may be glycinergic (see above). However, the presence of glycine receptors in granule cells is difficult to reconcile with the absence of any strychnine labelling in the cerebellum (Zarbin et al., 1981; see discussion Malosio et al., 1991). Gephyrin and/or the fl subunit are also produced by cells that do not express ~ subunits, such as the Purkinje cells. It has been postulated that gephyrin and/or the fl-subunit participate in other transmitter-gated ion channels, like the GABAAreceptors (see Malosio et al., 1991; Kirsch et al., 1993). For instance, Triller al. (1987) showed that gephyrin-immunoreactive postsynaptic specialisation face GABAergic synapses in the cerebellar cortex (Triller et al., 1987). Chen and Hillman (1993b) demonstrated that gephyrin-immunoreactive postsynaptic membrane specialisations (labelled with antibody R7A) faced a subpopulation of glutamate decarboxylase (GAD) immunoreactive boutons in the cerebellar nuclei. Since glycine often colocalizes with GABA it is possible that GABAergic boutons facing gephyrin-immunoreactive postsynaptic membrane specialisations, are also glycinergic. 3.8. M O N O A M I N E R G I C A F F E R E N T SYSTEMS AND RECEPTORS Several monoaminergic pathways terminate in the cerebellum (And6n and Ungerstedt, 1967). Their terminations in the cerebellar nuclei and the inferior olive will be considered elsewhere (Sections 5.2. and 6.3.2.2.). The monoaminergic fibers are primarily serotoninergic and noradrenergic (NA). The serotoninergic and noradrenergic fibers are present in all three layers of the cerebellar cortex. Their morphology differs from the classical mossy and climbing fiber afferents. Serotonin

Serotoninergic fibers in the cerebellar cortex were first demonstrated with the histofluorescence method, and were mainly recognized as transverse fibers in the molecular layer (H6kfelt and Fuxe, 1969). In studies using specific uptake, [3H]serotonin accumulated in small calibre, varicose axons. The varicosities were filled with minute, 15-25 nm round or flattened agranular vesicles and contained some large granular vesicles with a diameter of 50-90 nm (Beaudet and Sotelo, 1981). Synaptic contacts of the varicosities were few, most of them being present in the molecular layer. These, and other studies

A

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Fig. 74. Camera lucida drawing of serotonin immunostainingin Lobule X of the cerebellum of the rat (A) and

the paramedian lobule (PML) (B). Sagittal sections. Grl = granular layer; ML = molecularlayer; WM = white matter. Calibration bar = 200/lm. Bishop and Ho (1985). 102

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Fig. 75. Photomicrographs of serotonin immunoreactive fibers in selected regions of rat cerebellar cortex. A. Dark-field photomicrograph. B. High-power bright-field photomicrographs illustrating the beaded nature of the immunoreactive fibers (arrows). Vermal lobule X. Abbreviations: see Fig. 74. Calibration bar = 200 tim. Bishop and Ho (1985).

using specific uptake of [3H]serotonin also reported labelling over mossy fiber rosettes (Chan-Palay, 1975; Bloom et al., 1971). The presence of serotoninergic mossy fibers could, however, not be confirmed immunocytochemically with antibodies directed against conjugates of serotonin in rat, cat and opossum (Takeuchi et al., 1982; Bishop et al., 1985; Kerr and Bishop, 1991). Serotonin-like immunoreactivity was present in randomly oriented fibers in the molecular layer in the rat. Long transverse fibers were present in superficial strata of the molecular layer in the rat (Takeuchi et al., 1982) or in deep parts of the molecular layer in the opossum (Bishop et al., 1985), radially oriented fibers were most numerous in the vermis of rat (Figs 74 and 75) (Bishop and Ho, 1985). The granular layer contains a loose serotoninergic plexus. A layer of fine, beaded serotonin-containing fibers is present below the Purkinje cells in rat and opossum (Bishop and Ho, 1985; Bishop et al., 1985). The Purkinje cell layer only contains passing fibers. In the caudal vermis of the oppossum the serotoninergic innervation of the cortex is concentrated in midsagittal and parasagittal bands. In the cat serotonin-immunoreactive fibers are dense in the granular and Purkinje cell layers with only a few fibers in the molecular layer (Kerr and Bishop, 1991). These serotoninergic axons and varicosities have a uniform distribution throughout all lobules of the cerebellum with the exception of lobule X were the fiber density is low (Kerr and Bishop, 1991). A dense plexus of serotoninergic fibers is also found in all of the deep nuclei of the cat. In addition, there appear to be serotonin positive cell bodies in the deep nuclei (Kerr and Bishop, 1991) (Section 5.7.). Double-labelling experiments in rat (Bishop and Ho, 1985) (Fig. 76), opossum (Walker et al., 1988) and cat (Kerr and Bishop, 1991) revealed that the serotoninergic innervation of the cerebellum originates from the reticular formation. In the rat these neurons are distributed over the medullary and the pontine reticular formation (nucleus reticularis gigantocellularis, nucleus reticularis paragigantocellularis and nucleus pontis oralis). The origin of these fibers from the gigantocellular reticular formation and from a few neurons located in the medullary pyramids is more restricted in the opossum. Kerr and Bishop (1991) found that the serotoninergic projection to the cerebellar cortex of the cat shows some degree of topographical organization: Serotoninergic fibers in the anterior vermis arise from neurons located within the paramedian reticular nucleus, the 103

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lateral reticular nucleus and the lateral tegmental field, whereas fibers to the caudal vermis and the paramedian lobules exclusively derive from the lateral reticular nucleus. The hemispheres receive serotoninergic input from cells in the lateral tegmental field, the periolivary reticular formation and the paramedian reticular formation. Allthough retrogradely labelled cells were found in the raphe, in no case these cells were serotoninergic (Kerr and Bishop, 1991). Serotonin receptors

Physiological studies indicate that serotonin may play a modulatory role in the cerebellar circuitry (see Gardette and Cr6pel, 1993; Kerr and Bishop, 1992; and references therein). Serotonin interacts with a particularly large number of functionally and pharmacologically diverse receptor types. The serotonin receptors have been placed into five subgroups, 5HT1, 5HT 2, 5HT3, 5HT4 and 5HT5 (Tecott and Julius, 1993). The 5HT3 receptors are ionotropic receptors that mediate rapid excitatory responses in neurons, whereas the other serotonin receptor types are coupled to G-proteins, through which they activate second messenger cascades. The 5HT1 receptors comprise a heterogeneous group with at least six subtypes, named 5HT1A-5HT1F receptors. Ligand binding autoradiography with [3H]serotonin, which preferentially labels 5HT1 receptors, shows that a low level of [3H]serotonin binding occurs over the cerebellar cortex with the highest density over the molecular layer (Pazos and Palacios, 1985). The deep nuclei presented a higher density of [3H]serotonin binding with a lateral-to-medial decrease in receptor density. The cerebellar [3H]serotonin binding sites have the pharmacological properties of 5HT1B receptors (Pazos and Palacios, 1985). More recent autoradiographic studies with ligands specific for the 5HT1B and the 5HT1D receptors, which are closely related to 5HT1B receptors, indicate that the cerebellar cortex has 5HT1B, but no 5HT1D receptors (Bruinvels et al., 1993). The presence of 5HT1B receptors in rat cerebellum was confirmed by in situ hybridisation studies showing that 5HT1B receptor mRNA is expressed at high levels by Purkinje cells, but also by cells in the molecular layer, and in the deep cerebellar nuclei (Appel et al., 1990; Voigt et al., 1991; Maroteaux et al., 1992). In view of the findings that 5HT1B mRNA levels are relatively high in Purkinje cells, whereas receptor binding is low in the molecular and Purkinje cell layers, but relatively high in the deep nuclei, one is tempted to speculate that the 5HT1B receptor is localized presynaptically on Purkinje cell axon terminals. The ultrastructural localization of the 5HT1B receptors remains, however, to be determined. No 5HT1A receptors were found in the rat cerebellum with receptor autoradiography or in situ hybridisation (Pompeiano et al., 1992). Using immunocytochemistry with an antibody specific for the 5HT1A receptor, Matthiesen et al. (1993), however, found that the 5HT1A receptor occurs in Purkinje cells of immature, but not in adult cerebellum. The 5HTlc and 5HT 2 receptors were found at low concentrations in the deep nuclei in autoradiographic studies (Pazos et al., 1985; Molineaux et al., 1989). Since 5HT 2 and 5HTlc are structurally related they have been recently renamed as 5HTzA and 5HTzc receptors, respectively. In situ hybridisation studies have shown that mRNAs of both receptor types are expressed in the deep cerebellar nuclei (Pompeiano et al., 1994). The other known serotonin receptors (e.g. 5HT3 and 5HT4) seem to be essentially absent from rodent cerebellum (Tecott and Julius, 1993; Domenech et al., 1994). Summarizing, the known serotonin receptors are expressed at low levels or are not expressed at all in the cerebellum, with the exception of the deep nuclei which express significant levels of different types of serotonin receptors.

104

The cerebellum." chemoarchitecture and anatomy

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Fig. 76. Schematic drawings showing the distribution of labelled neurons in the medullary and pontine reticular formation in the rat, after injections of H R P in the vermis and adjacent parts of the hemisphere. Filled triangles represent neurons retrogradely labelled with H R P alone; closed circles represent neurons immunolabelled for serotonin alone; and asterisks represent double-labelled neurons. Each symbol represents one neuron in each population. Abbreviations: BC = brachium conjunctivum; BP = brachium pontis; CP = cerebral peduncle; D R = dorsal raphe nucleus; EC = external cuneate nucleus; ICP = inferior cerebellar peduncle; IN = interpeduncular nucleus; IOC = inferior olive; M R = nucleus raphe magnus; MV = medial vestibular nucleus; PG = nucleus paragigantocellularis; PH = nucleus pre-positus hypoglossi; PO = nucleus pontis oralis; R G = gigantocellular reticular nucleus; R M = nucleus raphe magnus; RO = nucleus raphe oralis; RP = nucleus raphe pallidus; SV = superior vestibular nucleus; V = nucleus of spinal tract of the trigeminal nerve; VII = nucleus of facial nerve. Redrawn from Bishop and Ho (1985).

Noradrenalin

The noradrenergic (NA) innervation of the cerebellum is mostly directed at the cortex. In the original studies of H6kfelt and Fuxe (1969) with a histochemical fluorescence method for the demonstration of catecholamines (Falck et al., 1962), NA containing fibers were found to be present as a randomly oriented plexus in all three layers of the cortex of rat cerebellum. The density of the NA innervation was lower in the flocculonodular lobe and the uvula than in the corpus cerebelli. A similar plexus was illustrated in rat cerebellum by Grzanna et al. (1989), using an antibody against a conjugate of NA (Geffard et al., 1986), and by Fritschy and Grzanna (1989) with an antibody against dopamine-fl-hydroxylase (Fig. 77A), the synthetic enzyme that converts dopamine to 105

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NA. A heavy plexiform innervation of the granular layer by thin varicose axons and parallel fiber-like fluorescence in the molecular layer was also found in chicken cerebellum. The parent fibers in the cerebellar white matter were concentrated in two parasagittal bundles (Mugnaini and Dahl, 1975). Axons containing dark core vesicles, which were labelled by specific uptake of [3H]NA terminated on Purkinje cell proximal dendrites and spines in the molecular layer (Bloom et al., 1971; Yamamoto et al., 1977). The inhibitory action of NA on Purkinje cells and the transduction of this response by cyclic AMP were reported in the subsequent papers of Hoffer et al. (1971) and Siggins et al. (1971). Purkinje cells therefore seem to be the main target of the NA innervation (Felten et al., 1986; see also Llano and Gerschenfeld, 1993). The ultrastructure of NA fibers of the cerebellar cortex and other parts of the rat CNS was analyzed with pre-embedding dopamine-fl-hydroxylase immunohistochemistry by Olschowka et al. (1981). Immunoreaction product was present in the axoplasm, associated with smooth endoplasmatic reticulum, Golgi apparatus, synaptic and large dense core vesicles and the outer membranes of mitochondria. Large varicosities were interconnected by narrow intervaricose axon segments. Varicosities, filled with clear, round synaptic vesicles and large dark-core vesicles, made asymmetric contacts with dendrites, but never with somata or axons. More than 50% of the labelled varicosities in the cerebellum made synaptic contacts: most of them with dendritic shafts, fewer on spines. Kimoto et al. (1981) used the histofluorescence method for light microscopy and potassium permanganate fixation in their electron microscopic studies of NA in rat cerebellum. They found NA-containing nerve terminals making contact with granule cell dendrites and secondary and spiny branchlets of the Purkinje cell dendritic tree. The terminals contained a large number of small dark core vesicles and a couple of larger ones. No synaptic contacts with the Purkinje cell somata were observed. Triarhoe and Ghetti (1986) are of the opinion that these results should be reconsidered because a distinction between NA and serotonin is not possible with this fixation method. The cerebellar NA fibers in the rat take their origin from the dorsal part of the locus coeruleus (group A6 of Dahlstr6m and Fuxe, 1964) and the A4 groups in the roof of the fourth ventricle. Single NA neurons innervate both the cerebellum and the forebrain (see also Steindler, 1981). The fibers enter the cerebellum close to the fourth ventricle. According to Pickel et al. (1973) the main entrance route is through the superior cerebellar peduncle. Pasquier et al. (1980) localized the cells of origin of the NAprojections to the cerebellum in the rat with specific retrograde transport of dopaminefl-hydroxylase. They traced the main projection from the entire caudal pole of the locus coeruleus and smaller contributions from the nucleus subcoeruleus and the cell groups A7 and A5; all projections are bilateral. The cerebellar projections in the cat take their origin from NA-containing cells located around the superior cerebellar peduncle, and the dorsolateral part of the locus coeruleus (Chu and Bloom, 1974; Somana and Walberg, 1978; see also Dietrichs, 1985, 1988). Grzanna et al. (1989), Fritschy and Grzanna (1989) and Grzanna and Fritschy (1991) reported a selective loss of the terminal portions of NA axons originating from the locus coeruleus in rats treated with the neurotoxin DSP (N-(2-chloroethyl-N-ethyl-2 bromobenzylamine)). DSP affects the NA innervation of the cerebellum, the cerebral cortex, the pontine nuclei, the inferior olive, the vestibular and cochlear nuclei, the sensory nuclei of the trigeminal nerve and the dorsal horn but spares the NA axonal plexus in the basal forebrain, the hypothalamus and most of the brain stem and the cord. After 24 hours loss of NA-staining is most-pronounced in the molecular layer, but with longer survival times most NA fibers have disappeared (Fig. 77B). 106

The cerebellum." chemoarchitecture and anatomy

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Fig. 77. Effects of DSP-4 (50 mg/kg) on the noradrenaline innervation of the cerebellar cortex of the rat. A. Control section incubated with an antibody against dopamine-fl-hydroxylase. B. Section from a DSP-4-treated rat. Parasagittal plane; dark-field photomicrographs. Magnification 90x. Fritschy and Grzanna (1989).

Adrenergic receptors Adrenergic receptors can be subdivided in ~l,0t2 and fll,fl2 receptors on the basis of p h a r m a c o l o g i c a l criteria (Bylund and Pritchard, 1983; M i n n e m a n et al., 1981). M o s t 107

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Fig. 78. A. x28 and B. x42 power photomicrographs of autoradiographs of ]2Siodocyanopindol (~2SlCYP) binding to adrenergic beta receptors in rat cerebellar cortex illustrating the high density of silver grains in the molecular layer and 'patches' of increased receptor density over the Purkinje cell layer, m, molecular layer; g, granule cell layer. (C) shows the patches at x 140 magnification with dark-field illumination. Sutin and Minneman (1985). (

receptors in the cerebellum are of the fl type, low levels of specific binding of the ~2 agonist [3H]para-aminoclonidine are associated with the granular layer in rat cerebellum (Unnerstall et al., 1984). Binding assays with the fl-adrenergic antagonist [3H]dihydroalprenolol in homogenates of cat cerebellum showed that the receptors are mainly of the f12 type and that they are evenly distributed over the vermis, the hemisphere and the cerebellar nuclei (Pompeiano et al., 1989). Elevated binding to the molecular layer was found in autoradiographs of tissue sections of rat cerebellum with ligands which bind non-selectively to fl receptors (Palacios and Kuhar, 1980). The same pattern emerged when either fll or f12 receptors were eliminated by specific inhibitors, with high levels of f12 binding in the molecular layer and much lower levels over the Purkinje and the granule cells. The overall contribution of fll receptor binding was low (Rainbow et al., 1984a). Beta adrenergic receptor binding was evenly distributed over the molecular layer, but at the level of the Purkinje cell somata the ligand is bound in irregular patches (Fig. 78) (Sutin and Minneman, 1985). These patches are most prominent in vermis and paravermal zones of the lobules I-IX and are less frequent in the medial hemisphere. Incubations with specific fll and f12 antagonists showed that the receptors in these patches are of both subtypes. 109

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Polyclonal antibodies which recognize domains of the f12 adrenergic receptor other than the NA-binding site, immunoreact with distal segments and dendritic spines of rat Purkinje cells. Immunoreactivity is present throughout the dendritic cytoplasm and accumulates at the dendritic membrane, opposite synaptic clefts (Strader et al., 1983). Dopamine

Dopaminergic projections to the cerebellum are still controversial. Low levels of dopamine (DA) supposedly represent a precursor of noradrenaline. DA-like immunoreactivity, as demonstrated with an antibody against a conjugate of DA, was present in fibers with irregularly spaced varicosities in all layers of the cerebellar cortex of the rat (Panagopoulos et al., 1991). Radially oriented DA fibers were abundant in the molecular layer. These fibers may take their origin from the A10 dopaminergic cell group of Dahlstr6m and Fuxe (1964) in the medial tegmental area. Cerebellar DA levels in rat cerebellum decreased by 50% after lesions of the A10 area (Kizer et al., 1976) and connections of the A10 area with the cerebellar nuclei and the granular and Purkinje cell layers were demonstrated with antegrade and retrograde axonal transport methods by Simon et al. (1979). Recently Ikai et al. (1992) reported a dopaminergic projection of the A10 group in the rat to restricted portions of the cerebellar hemisphere (Figs 79A,B and 80). Projections of the A10 group to the lateral cerebellar nucleus, however, appeared to be non-dopaminergic, as demonstrated immunocytochemically with an antibody against tyrosine hydroxylase, the enzyme involved in the conversion of tyrosine to DA. Fibers and terminals in the cortex were most numerous in the granule and Purkinje cell layer and scarce in the molecular layer. Their distribution differs, therefore, from the DA fibers described by Panagopoulos et al. (1991), which predominate in the molecular layer. Dopamine receptors

Five different receptors for dopamine have been distinguished, D~-Ds, which are often grouped as Dl-like (D1 and Ds) and Dz-like (D2, D3, and D4) receptors with high affinities for [3H]SCH23390 and spiperone, respectively (Grandy and Civelli, 1992). The D~ receptor is considered to be stimulatory, whereas the D 2 receptor is either inhibitory or unlinked to adenylate cyclase. Autoradiographic studies using [3H]SCH23390 as the ligand show that D~ receptor binding in the cerebellar cortex is low (Camps et al., 1990; Mansour et al., 1992). According to Camps et al. (1990), who compared the distribution of D~ and D 2 receptors in the cerebellum of rat, mouse, guinea pig, cat, monkey and man, Dl-binding was only detected in the Purkinje cell and molecular layer in rat and cat. Dz-receptors, labelled with [3H]CV 205-502, were only found at significant levels in rat cerebellum, specifically localized in the molecular layer of lobule X and IXc. In lobule IX high activity is distributed in sagittal and parasagittal columns. Although similar labelling has been repeatedly found with other 'D2-1igands' it was recently shown by both autoradiography and in situ hybridisation that the dopamine receptors in the rat nodulus and ventral uvula are of the D3-type. Thus Bouthenet et al. (1991) noticed a very low D 2 signal over the granular layer of all cerebellar lobules, and a high expression of D3 mRNA by Purkinje cells in lobule X and IXc. Autoradiographic studies with D3-preferring ligands, [3H]quinpirole (Gehlert, 1993; Levant et al., 1993) and 7-[3H]hydroxy-N,N-di-n-propyl-2-aminotetralin (L6vesque et al., 1992) showed a similar type of labelling in rat cerebellum (Fig. 81). The functional role of the cerebellar D3 receptor is puzzling in view of the lack of significant dopaminergic innervation. It is 110

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Fig. 79. Photomicrograph of camera lucida drawing (inset) of anterogradely labelled axons and terminal boutons in the crus I ansiform lobule of rat cerebellum after cholera toxin (b fragment) injection into the contralateral ventral tegmental area. Arrows point to 'rosettes' characteristic of mossy fiber endings in the granular layer (G). R Purkinje cell layer; WM, white matter. Scale bar = 40/xm. Ikai et al. (1992).

notable that substance P receptors have a very similar distribution in rat cerebellum (Nakaya et al., 1994). 3.9. H Y P O T H A L A M O CEREBELLAR CONNECTIONS AND HISTAMINERGIC PROJECTIONS Hypothalamo cerebellar fibers were demonstrated with different anterograde axonal transport techniques in squirrel monkey, cat and rat (Dietrichs, 1984; Dietrichs and Haines, 1985; Haines et al., 1984, 1985, 1986). Most fibers enter the cerebellum bilaterally with an ipsilateral dominance from the central grey, passing medial to the superior cerebellar peduncle. They enter the granular layer, where they branch. Fibers passing around the Purkinje cell somata enter the molecular layer, where most of them assume a longitudinal course in the long axis of the folia (Fig. 82). Few labelled mossy fibers and no labelled climbing fibers were seen. These 'multilayer' hypothalamo cerebellar fibers are more frequent in the cortex of the flocculus and the vermis, but are also present in parts of the hemisphere. Some fibers project to the cerebellar nuclei (Dietrichs and Haines, 1985; Haines et al., 1990). The projection takes its origin from cells in the posterior, lateral and dorsal hypothalamic areas, the lateral mammillary nucleus and the periventricular nucleus (Dietrichs and Zheng, 1984; Dietrichs, 1984; Haines and Dietrichs, 1984, 1987; Dietrichs et al., 1985b). Immunocytochemical studies have demonstrated the existence of histaminergic neurons, which are concentrated in the tuberomammillary nucleus of the posterior hypothalamus, and which give rise to fibers to almost all parts of the brain (reviewed by Wada 111

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et al., 1991). A low and low to moderate density of histamine-immunopositive fibers has been observed in rat (Inagaki et al., 1988) and guinea pig cerebellum (Airaksinen and Panula, 1988), respectively. Like the hypothalamo cerebellar fibers, the histaminergic fibers were sparsely distributed to all cortical layers, and were more frequent in the vermis and the flocculus than in other lobules. A moderate density of histaminergic fibers occurred in the cerebellar nuclei and the inferior olive. Three subtypes of histamine receptors have been pharmacologically identified in the brain: H1, H2 and H3 receptors. H1 receptors are coupled to phospholipase C, through which the inositol 1,4,5-trisphosphate-Ca 2+ and diacylglycerol-protein kinase C cascades are activated. Receptor autoradiographic studies with [3H]mepyramine and [125I]iodobolpyramine show that essentially no H~ receptors occur in the rat and human cerebellar cortex (Palacios et al., 1981b), whereas high densities are found over the molecular layer of guinea pig and mouse cerebellum (Palacios et al., 1981b; Rotter and Frostholm, 1986; Bouthenet et al., 1988). Moderate densities were found in mouse and guinea pig deep cerebellar nuclei (Rotter and Frostholm, 1986; Bouthenet et al., 1988). H1 receptor density over the molecular layer was greatly decreased in Purkinje cell deficient mice, whereas in reeler mice, which contain malpositioned Purkinje cells, high H~ receptor density was found over regions with heterotopically located Purkinje cells. No change in H1 receptor density was found in weaver mouse cerebellum, which is almost devoid of cerebellar granule cells. These data indicate that H~ receptors are predominantly localized on Purkinje cell dendrites (Rotter and Frostholm, 1986). 112

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Fig. 81. Bright-field autoradiographs showing [3H]quinpirole binding to dopamine D 3 receptors to coronal sections of the caudal cerebellum of rat. Note high-density of binding sites over the molecular layer (m) of the nodulus (X). Also note stripes of increased [3H]quinpirole binding over molecular layer of the uvula (IX). g, granular layer; IO, inferior olive; MVe-medial vestibular nucleus; 12, hypoglossal nucleus. Gehlert (1993).

H 2 receptors, which are coupled to adenylate cyclase were also found in the molecular layer of guinea pig cerebellum (Ruat et al., 1990). H3 receptors are autoreceptors involved in the inhibition of the release of histamine. A low level of H3 receptors were found to be homogeneously distributed throughout the rat cerebellar cortex (Arrang et al., 1987; Pollard et al., 1993). 3.10. CHOLINERGIC SYSTEMS AND ACETYLCHOLINESTERASE (ACHE) IN THE CEREBELLUM 3.10.1. Distribution of choline acetyltransferase Biochemical measurement of distinct levels of acetylcholine (McIntosh, 1941; Kfisa et al., 1982) and its biosynthetic enzyme, choline acetyltransferase (CHAT) in cerebellar tissue (K/tsa and Silver, 1969; Salvaterra and Foders, 1979; Hayashi, 1987; and others) indicated the presence of a cholinergic innervation in the cerebellum. ChAT activity varies among different lobules with the highest levels in the nodulus and ventral uvula. Following deafferentation of the cerebellar cortex, ChAT activity is considerably de113

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114

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creased, indicating that the cholinergic innervation is mostly of extracerebellar origin (K/tsa and Silver, 1969; Asin et al., 1984). Acetylcholine and ChAT were found in the glomerular fraction consistent with the view that acetylcholine may be the transmitter in a subpopulation of mossy fibers (Isr~iel and Whittaker, 1965; Bal/tzs et al., 1975). Early anatomical studies have employed acetylcholinesterase (ACHE) histochemistry. However, AChE activity is disproportionally high in the cerebellum (reviewed in Silver, 1974), and its widespread distribution in non-cholinergic cells and fiber systems precludes its use as a marker for cholinergic neurons. More reliable methods to unravel the anatomy of the cerebellar cholinergic system became available with the development of antibodies specific for CHAT. Early immunocytochemical studies revealed a subpopulation of ChAT-positive mossy fibers in rabbit and man (Kan et al., 1978, 1980). However, subsequent studies with monoclonal antibodies, failed to demonstrate ChAT positive staining in the cerebellum (Armstrong et al., 1983; Houser et al., 1983). Yet more recent studies by Ojima et al. (1989) in rat, by Illing (1990) in cat, by Barmack et al. (1992a) in rat, rabbit, cat, and monkey, and by DeLacalle et al. (1993) in man, demonstrated the presence of at least four types of cholinergic innervation in the cerebellum: (1) a subpopulation of ChAT-positive mossy fibers which primarily innervate the nodulus and the ventral uvula of the vermis; (2) a sparse plexus of thin beaded fibers which is present in all layers; (3) thin ChAT-positive fibers that innervate the cerebellar nuclei; and (4) a still controversial subpopulation of cholinergic Golgi cells. In addition, Ikeda et al. (1991) reported some typical granule cells with their bifurcating parallel fibers to be ChAT-immunoreactive in the cat, but this finding was not confirmed by others. Ojima et al. (1989) showed that a significant number of ChAT-positive mossy fiber rosettes was present throughout most vermal lobules, but that the density in the nodulus and ventral uvula was approximately ten times that of other vermal lobules (Fig. 85A). ChAT-positive mossy fiber rosettes were also enriched in the flocculus and the ventral paraflocculus. The distribution of ChAT-positive rosettes in rat observed by Barmack et al. (1992a) was essentially similar, although they suggested that the ChAT-positive rosettes were also enriched in lobules I, II and III (Fig. 83). They also demonstrated that ChAT-positive rosettes were concentrated in the nodulus and the ventral uvula of rabbit (Fig. 84), cat and monkey, and that the distribution of ChAT-positive profiles correlated with the distribution of ChAT activity. Large and small mossy fiber rosettes and transitional forms with the plexus of beaded fibers were distinguished in different species (Barmack et al., 1992a). Illing (1990) noted a 'wealth of ChAT-immunoreactive mossy fibers' that were slightly enriched in the ventral folia I to III, IX and X. ChAT-positive mossy fibers were also observed in human cerebellum, but their spatial distribution remains to be described (Kan et al., 1980; DeLacalle et al., 1993). By combining retrograde tracing and ChAT-immunocytochemistry Barmack et al. (1992b) demonstrated that the ChAT-positive mossy fibers in the nodulus, ventral uvula and the flocculus are likely to be secondary vestibular afferents. After injections of horseradish peroxidase in the lobules X and IX in rat and rabbit, neurons in the caudal medial vestibular nucleus and the nucleus prepositus hypoglossi were double labelled (Fig. 198). The flocculus and the paraflocculus of the rabbit receive a small, cholinergic projection, mainly from ChAT-positive neurons of the nucleus prepositus hypoglossi (Barmack et al., 1992b). Csillik et al. (1964) already drew the conclusion from the preferential distribution of AChE-positive glomeruli in the 'archi-cerebellum', that they represent terminals of a cholinergic system of secondary vestibulo cerebellar mossy fibers. Ikeda et al. (1991) noticed that in cat the number ChAT-positive mossy fibers 115

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Fig. 83. Choline-acetyltransferase (ChAT)-like immunoreactivity in the rat uvula-nodulus. A,B. Photomicrographs taken of lobule Xb. These photomicrographs demonstrate varieties of ChAT-positive mossy fiber rosettes in the granule cell layer and in the molecular layer of the rat nodulus. C. Illustration of ChAT-positive staining of mossy fiber rosettes and fine fibers innervating both the granule cell layer and molecular layer of the nodulus (lobule Xb). D. Illustration of three kinds of ChAT-positive mossy fiber rosettes found in the rat cerebellum: (1) Large grape-like terminals found with highest density in the uvula-nodulus (lobules IXb, Xa,b), (2) smaller mossy fiber rosettes that are characteristic of both the uvula (lobule IXa) and the anterior vermis (lobule I), (3) finely beaded ChAT-positive fibers that are found in both the granule cell layer and molecular layer throughout the cerebellum. Primary antibody: rat-~pig-ChAT. Barmack et al. (1992a). (

were reduced following excitotoxic and electrolytic lesions in the cerebellar nuclei (see below). The distribution of ChAT-positive mossy fibers roughly corresponds to that of unipolar brush cells (see Section 3.6.3.), which raises the question whether these mossy fibers innervate these cells. Electron microscopic analysis of ChAT-immunoreactivity in the nodulus showed that a minority (10-20%) of ChAT-immunoreactive mossy fiber terminals synapse on brush cell profiles, and that a minority (10-30%) of the mossy fiber terminals contact unipolar brush cells that are immunoreactive for ChAT (Jaarsma, 1995c). Barmack et al. (1992a) noted a dense population of small ChAT-positive mossy fiber-like terminals in the tip of lobules IXa,b of rat cerebellum that are significantly smaller than the other ChAT positive rosettes (Fig. 83). This type of labelling was not described by Ojima et al. (1989) but could be observed in their Fig. 4. The source and characteristics of this peculiar population of terminals is still unknown. Ojima et al. (1989), Illing (1990), Ikeda et al. (1991), Barmack et al. (1992a), and DeLacalle et al. (1993) observed thin ChAT-positive beaded fibers which were distinct from mossy fiber profiles. According to Ojima et al. (1989) these fibers in rat were most frequent immediately beneath or within the Purkinje cell layer. A substantial number of varicose fibers could also be identified in the molecular layer (Fig. 85B). Illing (1990) in cat noted that thin beaded fibers in the molecular layer mostly course at right angles from the Purkinje cell layer to the surface of the cerebellar cortex, whereas in the granular layer these fibers have an irregular course. The source of the sparse plexus of 'non-mossy' ChAT-immunoreactive fibers is presently unknown. It has been proposed that they originate in the ponto-mesencephalotegmental cholinergic complex (Illing, 1990), but at least for rat this seems to be untrue, since not a single retrogradely labelled, cholinergic neuron was found in the pontomesencephalic tegmentum after tracer injections in the cerebellar cortex (Woolf and Butcher, 1989). The cerebellar nuclei, instead, receive afferents from tegmental cholinergic neurons (see next paragraph). Ojima et al. (1989) showed that all cerebellar nuclei in rat are innervated by CHATimmunoreactive fibers. The density of these fibers varies between the different nuclei. Moderately dense innervation was found in most of the medial nucleus and in the magnocellular part of the lateral nucleus, whereas only a few ChAT-immunoreactive fibers invade the ventromedial parvicellular portion of the lateral nucleus and most of the interposed. Also in the cerebellar nuclei of man a moderate density of ChAT-positive fibers has been observed (DeLacalle et al., 1993). Both in rat and man these fibers did not form pericellular networks. DeLacalle et al. (1993) and Ikeda et al. (1991) also found 117

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sporadic ChAT-positive neurons in the cerebellar nuclei. As demonstrated by Woolf and Butcher (1989) with retrograde tracing, at least a portion of the cholinergic fibers in the cerebellar nuclei are likely to originate from the pedunculopontine tegmental nucleus. The projections are bilateral, but most prominently ipsilateral (Woolf and Butcher, 1989). Cerebellar Golgi cells display strong AChE activity (Shute and Lewis, 1965; Brown and Palay, 1972 and others), but there has been no evidence of ChAT-immunoreactive Golgi cells in the rat (Ojima et al., 1989; Barmack et al., 1992a; Jaarsma et al., 1995c). 118

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ChAT-positive Golgi cells have been found in cat (Illing, 1990; Ikeda et al., 1991) and in man (DeLacalle et al., 1993), although Barmack et al. (1992a) did not see them in their cat and monkey material. Illing (1990) found that the CHAT119

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Distribution and density of choline-acetyltransferase (ChAT)-immunoreactive Golgi cells (dots) in two subsequent 100 pm thick parasagittal sections through the vermis of the cerebellar of the cat. Roman numbers indicate lobules according to Larsell; pcs, pedunculus cerebellaris superior. Scale bar - 2 mm. B and C. Drawings of immunoreactive Golgi cells from 100 pm thick cerebellar sections. The uppermost cell is from the hemisphere, the lower one from the dorsal vermis. The arrows point to processes thought to be axons. ML, molecular layer; GL, granular layer. The border of gray to white matter is marked by a dashed line. Illing (1990). 120

The cerebellum." chemoarchitecture and anatomy

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positive Golgi cells mostly are positioned close to or in the Purkinje cell layer of both the vermal and hemispheral lobules (Fig 86). Also in man, ChAT-positive Golgi cells seem to be preferentially located beneath the Purkinje cell layer, but they were found to have a heterogeneous lobular distribution, with higher densities in the flocculonodular lobules (DeLacalle et al., 1993). In cat the ChAT positive Golgi cells have been estimated to represent only 5% of the total Golgi cell population (Illing, 1990). Ikeda et al. (1991) and DeLacalle et al. (1993) found ChAT immunoreactive neurons in the cerebellar nuclei in cat and man, respectively. According to Ikeda et al. (1991) cholinergic neurons in the cerebellar nuclei project to the cerebellar cortex in the form of mossy fibers, and to the thalamus and the red nucleus (see Section 5.2.).

3.10.2. Cholinergic receptors Nicotine receptors Both nicotine and muscarine receptors have been detected in the cerebellum. Nicotine receptors are cation-gating ion channel complexes and consist of five subunits, like the nicotine receptors in skeletal muscles. The subunit composition of neuronal nicotine receptor is found to be different from the muscle receptor, which is made up of four types of subunits (o~fle~) with two copies of the alpha subunit per receptor. A number of putative nicotine subunits are expressed in the mammalian brain, including at least five alpha subunits, named cz2, cz3, cz4, cz5, and cz7 and three subunits homologous to the muscle beta subunit, /32, /33 and/34 (Deneris et al., 1989; Duvoisin et al., 1989; Wada et al., 1989, 1990; Seguela et al., 1993). The subunits ~3, ~4, and/32 are expressed in rat cerebellum, the granular layer displaying low levels of cz3, cz4 and/32 mRNA, and Purkinje cells expressing high levels of/32 (Wada et al., 1989). Most Purkinje cells do not express any of the alpha subunits, but a few Purkinje cells display strong hybridisation signals with the cz2, ~3 or ~4 probes. Cells in the cerebellar nuclei express moderate levels of cz4 and/32 mRNA. A recent study with antibodies against the/32 subunit showed that the intensity of fl2-immunostaining was high in the perikarya and main dendritic arbors of Purkinje cells, low in granule cells and moderate in neurons of the cerebellar nuclei, thus confirming the results from in situ hybridisation (Hill et al., 1993). Swanson et al. (1987) using a iodinated monoclonal antibody to chicken neuronal acetylcholine receptors ([125I],mAB270) that is assumed to specifically immunoreact with the fl2-subunit of rat, obtained a somewhat different immunodistribution offl2-subunits, with labelling concentrated in the granular layer. Functional channels gated by nicotinic agonists are formed by certain combinations of alpha and beta subunits (Deneris et al., 1991). Receptors made of cz4 and f12 subunits are most common in the brain, and are believed to represent high-affinity [3H]acetylcholine binding sites, (that can be labelled in the presence of atropine to block muscarinic sites), and the [3H]nicotine binding sites of the brain (Clarke, 1993). Accordingly in rat, high-affinity [3H]nicotine binding sites are present at low density in the granular layer and the cerebellar nuclei, which are the structures expressing both cz4 and //2 subunits, but are essentially absent in the Purkinje cell and molecular layer, where ~4 subunits are lacking (Clarke et al., 1985). It should be noted that Purkinje cells, in spite of the fact that they express high levels of fl2 subunit, may lack functional nicotine receptors, because of the absence (as far as known) of alpha subunits. The cz7 subunit is different from the other neuronal nicotine receptor subunits in forming functional channels in the homomeric configuration (S6guala et al., 1993) that, unlike other nicotine receptors of the brain, are sensitive to ~-bungarotoxin (czBTX).

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The distribution of ~7-mRNA throughout the brain roughly correspond to that of [125I]~BTX binding sites. It is therefore assumed that nicotine receptors made up of ~7 subunits represent the major portion of brain ~BTX binding sites (Clarke et al., 1985; S6gu61a et al., 1993). The level of [125I]~BTX binding in most of the rat cerebellum is low. Patches of intense labelling were, however, found over glomeruli in the lobules I, IXd, X and the flocculus (Fig. 87; Hunt and Schmidt, 1978; Frostholm and Rotter, 1986). Frostholm and Rotter (1986) postulated that these [125I]0~BTX binding sites were located presynaptically on mossy fibers, based on the observation that developmental 122

The cerebellum." chemoarchitecture and anatomy

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appearance of the binding sites corresponds to the appearance of the mossy fibers. Notably, the distribution of [125I]~BTX binding sites is similar to that of ChAT-positive mossy fibers rosettes, which raises the possibility that cerebellar '~BTX-receptors' are localized presynaptically on the subpopulation of cholinergic mossy fibers. The distribution of ~7 mRNA has yet to be studied systematically in the vestibulo-cerebellum, but data from in situ hybridisation presently available suggest that no ~7-mRNA is expressed in the cerebellum, which is consistent with the idea of a presynaptic localisation of '~7-receptors' (S6gu61a et al., 1993). Recently an immunocytochemical study on the distribution of the ~7 protein has been reported, in which three different monoclonal antibodies raised against chicken ~7 were employed (Dominguez del Toro et al., 1994). The results of this study, however, are puzzling since immunolabelling in the cerebellar cortex was mainly localized in Purkinje cells, which disagrees both with the distribution of ~7-mRNA and [125I]~BTX binding sites. Muscarine receptors

Muscarine receptors are coupled to G-proteins and may activate various second messenger cascades. At least four subtypes (M1, M2, M3 and M4) could be distinguished according to pharmacological criteria (e.g. see Waelbroeck et al., 1990), whereas molecular cloning has revealed five distinct subtypes (ml-m5; Hulme et al., 1990). Membrane binding and autoradiographic studies have shown that cerebellar muscarine receptors have the pharmacological properties of the cardiac (M2)-type (Mash and Potter, 1986; Spencer et al., 1986; Waelbroeck et al., 1987, 1990; Araujo et al., 1991; Aubert et al., 1992). Concordantly, the cerebellum has been demonstrated to primarily express m2-mRNA (Vilar6 et al., 1992, 1993) and m2 receptor protein (Levey et al., 1991; Li et al., 1991). It has been proposed that also m3 (respectively M3) receptors may be expressed in the cerebellum, because granule cells in culture express m3 mRNA (Fukamauchi et al., 1991). Immunochemical and pharmacological studies, however, indicate that m3 receptors represent less than 10% of the cerebellar muscarine receptor population (Waelbroeck et al., 1990; Wall et al., 1991). Moreover, no high-affinity binding sites for [3H]4-DAMP, a ligand with preference for M3 receptors, were found in the cerebellum (Araujo et al., 1991). Early autoradiographic studies on the topographic distribution of muscarine receptors in rat cerebellum with [3H] propylbenzilylcholine mustard as the ligand showed that muscarine receptors were confined to the molecular layer of the nodulus and the ventral uvula (Rotter et al., 1979a,b). Subsequent autoradiographic studies in rat with the more potent antagonist [3H]quinuclinidyl benzylate ([3H]QNB, Neustadt et al., 1988) and with M2-specific ligands [3H]oxotremorine (Spencer et al., 1986; Vilar6 et al., 1992), and [3H]AF-DX384 (Aubert et al., 1992), all gave identical results. Muscarine receptors were found to be localized throughout the whole cerebellar cortex, with the highest densities in the molecular layer of the nodulus and the ventral uvula (Fig. 88). The amount of muscarine receptors in the other lobules was relatively low, with higher densities in the granular and Purkinje cell layers than in the molecular layer (Fig. 88). Muscarine receptors also occur in the cerebellar nuclei (Spencer et al., 1986; Neustadt et al., 1988; Vilar6 et al., 1992) and in some parts of the white matter. In the caudal folia of the uvula (lobule IXb) muscarine receptors appear to be distributed in five parasagittal columns of high receptor density that traverse the granular layer (Fig. 88D, Neustadt et al., 1988). In situ hybridisation shows that m2 mRNA in rat cerebellum is localized in the granular layer as well as in the deep nuclei (Vilar6 et al., 1992). In accordance with the autoradiographic data the highest signal is present in the nodulus and ventral uvula. 123

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J. Voogd, D. Jaarsma and E. Marani

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Fig. 88. Bright-field photograph showing cresyl-violet staining (A and C) and corresponding dark-field photomicrographs showing the distribution of muscarine receptors using [3H]QNB binding (B and D, respectively) in the caudal cerebellum of the rat. A and B. Coronal section through the nodulus (X) and the rostral uvula (IX) showing high density of [3H]QNB binding over the molecular layer (m) of lobule X and ventral region of lobule IX. Also note higher binding densities at the level of the Purkinje cell layer (P) in most lobules. C and D. Coronal section at the level of the dorso-caudal uvula (IXa,b). Note five parasagittal columns (large arrows in D) of high receptor density that traverse the granular layer (g); w, white matter. Scale bars = 1 mm. Neustadt et al. (1988).

Since in this region the mRNA signal is in the granule cell layer, whereas receptor binding is high over the molecular layer (Fig. 89), one is tempted to speculate that the muscarine receptors are associated with parallel fibers. In fact, analysis of sections immunostained by Levey et al. (1991) with a polyclonal antibody specific for the m2 receptor, showed that parallel fibers are immunostained (Jaarsma et al., 1995a). Parallel fiber staining was seen in all lobules. It remains to be determined whether the amount of parallel fiber staining is most prominent in the nodulus and ventral uvula, concordant with the density distribution of receptor binding. In addition to parallel fibers the m2-antibody also immunostained Golgi cells and some mossy fiber rosettes (Jaarsma et al., 1995a). Neustadt et al. (1988) using receptor autoradiography noted that the topographic distribution of muscarine receptors in rabbit and guinea pig cerebellum is characterized by the presence of parasagittal columns of very high receptor density over the molecular layer (Fig. 90). In rabbit the bands of high muscarine receptor density were most prominent in the anterior lobe (lobules I-V), in Crus I and II, and in the flocculus and the ventral paraflocculus. Their distribution only partially corresponded to the Zebrinpositive zones (Jaarsma et al., 1995a). The receptors in the parasagittal bands were 124

The cerebellum." chemoarchitecture and anatomy

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Fig. 89. Comparison between the distribution of m2 muscarine receptor mRNA (A) and [3H]oxotremorine-M binding to muscarine M2 receptor sites (B) in approximately equivalent coronal sections of rat brain at the level of the caudal cerebellum. Note that m2 mRNAs in the ventral uvula (IX) and the nodulus are localized in the granular layer, whereas M2 receptor binding sites are in the molecular layer (m). 12, hypoglossalnucleus. Scale bar - 3 ram. Vilar6 et al. (1992).

located in Purkinje cell dendrites as demonstrated immunocytochemically with a monoclonal antibody specific for the m2 receptor protein. A second characteristic feature of the distribution of rnuscarine receptor in rabbit cerebellum, is the presence of a stripe of very high receptor density immediately above the Purkinje cell layer in the nodulus and the ventral uvula (Fig. 90B) (Neustadt et al., 1988). Immunocytochemistry showed that the receptors in this transverse stripe were localized on a population of densely packed parallel fibers (Jaarsma et al., 1995a). In addition to these parallel fibers and Purkinje cell dendrites, also Golgi cells, that in rabbit in part are located in the molecular layer (Spa~ek et al., 1973), and a subpopulation of mossy fiber rosettes are immunostained with m2-receptor antibody (Jaarsma et al., 1995a). Autoradiographic data from human (Cortes et al., 1987) and cat cerebellum (D. Jaarsma, unpublished observations), and immunocytochemical data from monkey (Jaarsma and Levey, unpublished) indicate that the distribution of muscarine receptors in the cerebellum of these species is different from that in rodents and rabbit. The level of ligand binding was very low with slightly higher receptor densities over the granular layer. There is no increased receptor density in the nodulus and the ventral uvula. The overall distribution in monkey was similar. Only Golgi cells and a subpopulation of mossy fibers rosettes, (mainly located in the vermal lobules III-VI), were immunostained (Jaarsma and Levey, unpublished). Summarizing it appears that the cellular and regional distribution of muscarine receptors in the cerebellum is different between different species. Golgi cells and subpopulations of mossy fibers seem to express muscarine receptors most constantly. An interesting aspect about the presence of muscarine receptors in parallel fibers in rat and rabbit, is that the lobular distribution of m2-containing parallel fibers, is the same as that of ChAT-positive mossy fiber rosettes (see above). This raises the possibility that muscarine m2 receptor are specifically expressed by those granule cells that are innervated by cholinergic mossy fibers. If this proves to be true, this would imply that there 125

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F Fig. 90. Bright-field photograph showing cresyl-violet staining (A, C and E) and corresponding dark-field photomicrographs showing the distribution of muscarine receptors using [3H]QNB binding (B, D and F, respectively) in the cerebellar cortex of the rabbit. A and B: coronal section through the nodulus (X) and ventral uvula (IX); C and D: coronal section through lobules of the rostral vermis. E and F: coronal section through the ventral paraflocculus and flocculus. Note columns of high [3H]QNB binding density in the molecular layer (m, arrows in B and F) and the stripe of high binding density at the level of the Purkinje cell layer (P) of the nodulus, the ventral uvula (B) and the a portion of the flocculus (F). g, granular layer; w, white matter. Scale bars = 1 mm. Neustadt et al. (1988).

is some regulatory interaction between the ChAT-positive mossy fibers and granule cells through which the expression of muscarine m2-receptor is regulated. Although lightmicroscopic analysis suggest that the muscarinic receptors are absent in granule cell dendrites at the synapses with ChAT-positive mossy fibers, this possibility can not be excluded without electron microscopic examination. The function of muscarine receptors in the parallel fibers is puzzling since there seems to be little cholinergic innervation in the molecular layer (see above). 126

The cerebellum." chemoarchitecture and anatomy

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Physiological responses to acetylcholine Only a few physiological studies on the actions of acetylcholine in the cerebellar cortex have been reported and the results of these studies were often contradictory: Iontophoretic application of acetylcholine has been observed to mildly excite Purkinje cells via a muscarinic (Crawford et al., 1966) or a nicotic (McCance and Phillis, 1968) mechanism, or to inhibit them via a mixed muscarinic and nicotinic mechanism. For interneurons no effect of acetylcholine was found by Crawford et al. (1966), whereas De la Garza et al. (1987) found a strong nicotinic excitatory effect. Cr6pel and Dhanjal (1982) reported slow depolarisations of Purkinje cells, accompanied by an increase in membrane resistance, with application of high doses of acetylcholine in slices of the nodulus and the ventral uvula of the rat's cerebellum, that was mediated through muscarine receptors.

3.10.3. Acetylcholinesterase Based on their substrate specificity cholinesterases can be subdivided into the acetylcholinesterases (AChE's) and the butyryl or pseudocholinesterases. Several isoenzymes of AChE have been detected in brain tissue. Three to four isoenzymes of AChE can be distinguished in developing brain (Henderson, 1977). In the chicken the faster migrating forms predominate in embryos. Their composition changes progressively into the slowly migrating forms of the adult tissues. Pseudocholinesterase can be distinguished from the acetylcholinesterases by specific inhibitors (iso-OMPA or ethopropazine) and by using butyrylcholiniodide as a substrate, which is split predominantly, though not exclusively by pseudocholinesterase. Most other esterases can be inhibited using eserine (Marani et al., 1977). Butyrylcholinesterase has also been localized with immunocytochemical methods (Barth and Ghandour, 1983). AChE-staining in the molecular layer of the mammalian cerebellum is generally lower than in the granular layer, but great variations occur and the pattern is reversed in the human and the avian cerebellum (Friede and Fleming, 1964). AChE-staining generally is higher in the vestibulocerebellum than in other parts of the cerebellum. The reactivity in the glomeruli in rat (Csillik et al., 1964; Brown and Palay, 1972 and many others) and rhesus monkey (Robertson and Roman, 1989) follows this general pattern, but the distribution of AChE-positive Golgi cells is more uniform (Brown and Palay, 1972). In the cerebellar white matter of monkey cerebellum AChE-rich fibers are distributed in parasagittal compartments, that are aligned with concentrations of strongly ACHEreactive regions in the granule cell layer and narrower, AChE-rich 'spikes' in the molecular layer (Hess and Voogd, 1986; Marani, 1986). Purkinje cells in certain parts of rat and guinea pig cerebellum, including the lobules IX and X of the caudal vermis, display a transient reactivity for ACHE, which disappears later. AChE was localized in adult Purkinje cells of the lobules IX and X (Robertson et al., 1991); these cells are arranged in multiple, sagittal bands (Gorenstein et al., 1987). Robertson et al. (1991), however, were unable to confirm the transient staining with AChE in rat Purkinje cells. AChE-positive displaced Golgi cells (Ramon y Cajal, 1911) are present in the lower one third of the entire molecular layer of the rabbit cerebellum (Spa~ek et al., 1973) and strong AChE-staining is present in this stratum in the vermis and certain lobules of the hemisphere in the same species (Tan et al., 1995a). An AChE-band pattern was detected in the molecular layer of the vermis of the anterior and posterior lobes in 2-4 month old cats (Marani and Voogd, 1977) (Fig. 120). 127

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J. Voogd, D. Jaarsma and E. Marani

Reaction product in transverse sections was present in narrow striations reaching from the AChE-negative Purkinje cell somata to the pial surface; in sagittal sections the distribution of AChE was more uniform. The same banding pattern could be visualized in adult cats, with AChE-histochemistry on aldehyde-fixed tissue (Brown and Graybiel, 1983). No such banding was present in the cerebellum of rats or primates (Marani, 1986). Spikes of strong AChE-activity sometimes extend from heavily stained portions of the granular layer into the molecular layer, at certain preferred localizations (Hess and Voogd, 1986). The ultrastructural localisation of AChE has methodological problems, because of uncontrolled diffusion of reaction product (e.g. see discussion Brown and Palay, 1972). In the molecular layer AChE-reaction product was located in subsurface cisterns of Purkinje cell dendrites and in the spine apparatus of their dendritic spines and, extracellularly, associated with the dendritic plasma membrane and around parallel fibers (Marani, 1982a, 1986). AChE was never found in the synaptic cleft of parallel fiberPurkinje cell synapses, lntra-axonal deposits in parallel fibers were considered as an artefact. Neurons in deep parts of the molecular layer that may represent basket or stellate cells, contain AChE-reaction product in their rough endoplasmatic reticulum. AChE in cat molecular layer, therefore, is associated with neurons and not with glial cells. AChE in the molecular layer of the rat is preferentially located in Bergmann glia (Friede and Fleming, 1964; Barth and Ghandour, 1983; Gorenstein et al., 1987; Robertson et al., 1991). Brown (1985a) demonstrated an increase in AChE-staining in the cat molecular layer, 2-6 weeks after lesions of the inferior olive. Upregulation of AChE was found in the projection zones of the lesioned olivary neurons. These plastic changes are reminiscent of the re-appearance of neonatal AChE in Purkinje cells after the transection of their axons (Phillis, 1968). AChE-staining in the granular layer is distributed in 'An alternating fine mosaic of stained and unstained spots, with regional differences in staining intensity' that represent AChE-positive Golgi cells and moderately to strongly stained glomeruli. Granule cell somata remain unstained in the rat cerebellar cortex (Altman and Das, 1970). The presence of AChE in glomeruli has been generally acknowledged (Gerebtzoff, 1959; Csillik et al., 1964; Shute and Lewis, 1965). AChE-reaction product in rat cerebellum is located outside the mossy fiber terminal, between the axonal plasma membrane and the dendrites of the granule cells and the Golgi cell terminals. AChE in Golgi cells is associated with the cisterns of the rough endoplasmatic reticulum and the perinuclear cisterns. Granule cells, Purkinje cells and glia are AChE-negative (Brown and Palay, 1972). The preferential staining of the borders of white matter compartments (the 'raphes', Voogd, 1964) for AChE (Hess and Voogd, 1986; Marani, 1986; Voogd, 1995; Fig. 126) and its use in topographical analysis of the corticonuclear and olivocerebellar projections will be considered in Sections 6.1.1., 6.1.2., 6.1.5. and 6.3.3. Several authors have attempted to devise other functions for AChE than the hydrolytic cleavage of acetylcholine; their proposals were reviewed by Greenfield (1984) and Appleyard and Jahnsen (1992). 3.11. NEUROGLIA The morphology of the neuroglia in the cerebellar cortex was reviewed by Palay and Chan-Palay (1974) and Ghandour et al. (1980). Three forms were distinguished, the Bergmann glial fibers in the molecular layer that take their origin from Golgi epithelial cells, located in the Purkinje cell layer, the astroglia and the oligodendroglia. The 128

The cerebellum." chemoarchitecture and anatomy

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Fig. 91. Bergmann glial fibers in the rat cerebella of various ages. PAP immunocytochemistry with anti-GFAP antiserum. The ages of the rats are 3, 5, 8 days and 30 months in A, B, C and D, respectively. EG, external granule cell layer; M, molecular layer; G, granule cell layer. Scale bars = 20/lm. De Blas and Cherwinski (1985).

enzyme histochemistry of cerebellar glia was studied by Sotelo (1967). Neuroglia has its own metabolism, that is qualitatively different from that of neurons. The glial anaerobic glycolysis is high in neuronal neuroglia and interfascicular neuroglia. The aerobic glycolysis is secondary and much weaker than in nerve cells. The hexose129

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J. Voogd, D. Jaarsma and E. Marani

Fig. 92. Micrograph of adult cerebellar cortex, incubation with vimentin antibody V9 counterstaining with cresylviolet. Sagittal unfixed cryostat section. Bergmann glial fibers (arrowheads) and large astrocytes (arrows) are seen in the molecular and granular layer, respectively, pc, Purkinje cell; ml, molecular layer; gl, granular layer. Bar = 100 pm. Roeling and Feirabend (1988)

monophosphate shunt is also important in glia. In oligodendroglia and astroglia different enzyme patterns were found. The main difference concerns glycogen, which is metabolised almost exclusively in astroglia (Sotelo 1967). Bergmann glia and astroglia are immunoreactive for anti-glial fibrillary acidic protein (GFAP) (Fig. 91). The development and the adult configuration of the Bergmann glia has been studied, using antibodies against GFAP (Bignami and Dahl, 1974; De Blas, 1984; De Blas and Cherwinsky, 1985; Levitt and Rakic, 1980; Gr~iber and Kreutzberg, 1985; Pelc et al., 1986). Bergmann glia can also be demonstrated with anti-vimentin (Dupouey et al., 1985; Roeling and Feirabend, 1988) (Fig. 92) and with cell-specific monoclonal antibodies (De Blas, 1984; De Blas and Cherwinsky, 1985; Edwards et al., 1986). Butyrylcholinesterase has been localized in Bergmann glia and in glial cells in the granular layer using an immunocytochemical method in the rat (Barth and Ghandour, 1983). Carbonic anhydrase is a specific marker for oligodendrocytes (Cammer, 1984). The distribution of oligodendrocytes in the cerebellar cortex of the mouse was studied with immunostaining for carbonic anhydrase II by Ghandour et al. (1980, 1981). Occasionally oligodendrocytes are present in the molecular layer, they are more common in the granular layer and abundant in the white matter, sn-Glycerol-3-phosphate-dehydrogenase (GPDH) in mice is present in adult oligodendrocytes (De Vellis et al., 1977), but also in Bergmann glia (Fisher et al., 1981) (Fig. 93). The expression of GPDH in Bergmann fibers is dependent on the adjoining Purkinje cells (Fisher, 1983). A similar interdependency may exist for 5'-N in Bergmann glia (Hess and Hess, 1986) (see Section 130

The cerebellum." chemoarchitecture and anatomy

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Fig. 93. Sagittal section through the cerebellum of a 25-day-old BALB/cBy mouse, with anti-sn-glycerol-3 phosphate dehydrogenase (GPDH) serum (1:10,000) viewed with a dark-field condenser. The molecular layer is densely stained although clear areas representing non-staining cell bodies and processes are numerous. Intensely stained cell bodies of the Bergmann glia are located between large unstained Purkinje cell somata, at the boundary between the molecular and granular layers. Magnification x l00. Fisher et al. (1981). 3.5.), for t a u r i n e t h a t shifts f r o m P u r k i n j e cells to B e r g m a n n glia u n d e r c o n d i t i o n s of h y p o - o s m o t i c stress ( N a g e l h u s et al., 1993) (see Section 3.1.2.) a n d Z e b r i n I, t h a t a p p e a r s in B e r g m a n n glia after lesions of the cerebellum ( D u s a r t et al., 1994).

Fig. 94. A and B. Distribution of 3-fucosyl-N-acetyl-lactosamine (FAL)-immunoreactive Bergmann glial cells in adult mouse cerebellum. Note zonal distribution of immunoreactive neuroglia in molecular layer (MOL) in A. GCL, granular layer. Bars in A = 800/~m, in B = 100 ~tm. Courtesy of Dr. J.K. Mai, Department of Neuro-anatomy, Heinrich Heine University, Dfisseldorf. 131

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J. Voogd, D. Jaarsma and E. Marani

Fig. 95. Chicken cerebellum, 1 day old, 1-/lm thick sections. Post-embedding immunocytochemistry with antibody IX-50 against a putative kainate receptor shows immunoreactivity outlining Purkinje cells (Pc), their main ascending dendrites (arrows) as well as their spiny branchlets. Immunoreactivity is associated with Bergmann glial cells (BG) which lie beneath and amongst the Purkinje cells. Scale bar = 20/~m. Somogyi et al. (1990).

Several substances are distributed in zonally distributed Bergmann glial cells. This may be the case for 5'-N, and has been observed for 3-fucosyl-acetyl-lactosamine (FAL) immunoreactivity in mouse cerebellum (Fig. 94) (Bartsch and Mai, 1991; Marani and Mai, 1992). The protein kinase types ~, flII, y and ( have been localized in Bergmann glia (Shimohama et al., 1990; Hidaka et al., 1988; Wetsel et al., 1992). Bergmann glial cells 132

The cerebellum." chemoarchitecture and anatomy

Ch. I

contain both nitric oxide synthase and guanylcyclase and are the main cerebellar store for cyclic GMP (see Section 3.4.). As mentioned before also homocysteic acid (Cu6nod et al., 1990; Grandes et al., 1991; Tschopp et al., 1992) (Figs 44 and 45) and 5'nucleotidase (Kreutzberg et al., 1978 and Fig. 59; see, however, Marani 1986 and Fig. 58) are associated with Bergmann glia. Climbing fiber-induced release of homocysteic acid and adenosine, therefore, involve this glial compartment (see Sections 3.2.2. and 3.5.). Bergmann glial cells and granular layer astroglia are provided with glutamate receptors and, therefore, may be actively involved in the cerebellar neural transmission (Fig. 95) (see Section 3.3.1.).

4. GROSS ANATOMY OF THE MAMMALIAN CEREBELLUM The gross anatomy of the cerebellum is the morphology of its cortical sheet. The cerebellum in mammals, birds and some reptiles is subdivided by transverse fissures of varying depth in lobes, lobules and folia. Two paramedian sulci demarcate the vermis from the hemispheres in the mammalian cerebellum. The present nomenclature for the mammalian cerebellum evolved from older purely descriptive studies of the human cerebellum (reviewed by Glickstein, 1987), elaborate comparative anatomical studies of the adult cerebellum (Elliot Smith, 1903; Bolk, 1906; Riley, 1928, reviewed by Larsell 1970; Larsell and Jansen, 1972) and comparative embryological investigations of the development of the folial pattern (Stroud, 1895; Bradley 1903, 1904; Larsell, 1947, 1952, 1953, 1954, 1970). The cerebellum of the rat served as the prototype for Larsell's subdivision of the mammalian cerebellum in 10 lobules (Larsell, 1952, 1970) (Fig. 96). HEMISPHERE

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Larsell (1952) and comparativeanatomical nomenclatureof Bolk (1906). Right. Nomenclatureof the human cerebellum. 133

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Fig. 97. Cerebellum of the rat. a. Anterior aspect with anterior lobe (left), without anterior lobe (right). b. Caudal aspect. c. Ventral aspect. d. Dorsal aspect. e. Topology of the caudal segments (paraflocculus and flocculus) of the folial chain of the hemisphere. Areas without cortex where the white matter comes to the surface, are indicated with light hatching; double hatching indicates cross-sectional area of cerebellar peduncles, solid black indicates roof of the fourth ventricle in c. ApmF = ansoparamedian fissure; CI(1I) = crus I(I1) of the ansiform lobule; Cop = copula pyramidis; FL = flocculus; IcF = intrecrural fissure; IPFLS = interparafloccular sulcus; mcb = middle cerebellar peduncle; PFL = paraflocculus; PflS = parafloccular sulcus; PLF = posterolateral fissure; PM = paramedian lobule; PmS = paramedian sulcus; PpF = prepyramidal fissure; PreculF = preculminate fissure; PrF = prmary fissure; PsF = posterior superior fissure; rb = restiform body; scp = superior cerebellar peduncle; SecF = secondary fissure; Sim = simple lobule. Modified from Voogd (1995).

The cerebellum." chemoarchitecture and anatomy

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Larsell extensively studied its adult morphology and the development of its folial pattern. More recent accounts of the cerebellum of the rat (Voogd, 1995) (Fig. 97) and the mouse (Marani and Voogd, 1979) are available. The cerebellum of the cat was described by Larsell (1953, 1970) and Voogd (1964) (Fig. 98). The gross morphology of the cerebellum in different species of macaques was described by Larsell (1953, 1970), and by Madigan and Carpenter (1971) for Macaca rhesus. The anatomy of the cerebellum of macaques is very similar to other primates (Macaca fascicularis, Fig. 99) (Larsell, 1970: Cercocebus sp. Cebus sp., Ateles, Saimiri sciurus) and subprimates (Haines, 1969). It is possible to recognize the main subdivisions in nearly all mammalian cerebella by inspection of the branching pattern of the arbor vitae in a midsagittal section of the vermis (Fig. 98) and the characteristic convolutions of the folia of the hemisphere. For the lobules of the vermis either the classical nomenclature of the human cerebellum or Larsell's (1952, 1970) numbering system can be applied. The anterior lobe consists of lingula (lobule I), the central lobule (II, III), and the culmen (IV, V). The base of the lingula is continuous with or embedded in the anterior medullary velum. The subdivision of the anterior lobe by the preculminate fissure into the lobules I-III and the lobules IV-V is of prime relevance to its connectivity. It is usually deep and its walls are subfoliated. The anterior lobe is separated from the posterior lobe by the deep primary fissure. The posterior lobe vermis can be subdivided in most mammals in the declive (VI), folium and tuber (VII A and VII B), the pyramis (VIII), the uvula (IX) and the nodule (X). The secondary and posterolateral fissures in between the lobules VIII, IX and X usually are quite distinct. The prepyramidal fissure separating lobule VII and VIII is less obvious in some species and the borders between the lobules VI, VII A and VII B often are difficult to recognize. For the lobules of the hemisphere of the mammalian cerebellum both Bolk's (1906) descriptive terms and Larsell's numerals can be used. In Larsell's nomenclature the hemispheral lobules bear de prefix H to the number of the vermal lobule with which they are continuous. Bolk's (1906) and Larsell's (1952, 1970) nomenclature for the hemisphere are not readily interchangeable because they are based on conflicting views on the morphology of the mammalian cerebellum (Fig. 96). Bolk (1906; see Glickstein and Voogd, 1995, for a recent discussion of Bolk's views) stressed the difference between vermis and hemisphere that should be considered as independent folial chains, which have a tendency to local longitudinal expansion. For the cat the folial chains are illustrated in Fig. 98. In the anterior lobe the folial chains of vermis and hemisphere develop in parallel. The inter- and most of the intralobular fissures continue uninterruptedly from the vermis into the hemispheres and a paramedian sulcus is either absent or shallow. The same situation prevails in the region of the posterior lobe immediately behind the primary fissure. Bolk coined the name 'lobulus simplex' to express the continuity of vermis and hemisphere in this part of the cerebellum. The vermal portion of the simple lobule corresponds to the declive (Larsell's lobule VI). Caudal to the lobulus simplex the folial chain of the hemisphere deviates from the vermis and forms a series of loops. The first loop is the ansiform lobule, that can be subdivided in the Crus I and the Crus II. The cortex in the center of the ansiform lobule, in between folium vermis (lobule VIIA of Larsell) and the hemisphere, usually is interrupted (Figs 97 and 99). This interruption may involve all layers of the cortex, with white matter coming to the surface, or only affect the parallel fibers of the molecular layer. At the level of the paramedian lobule the folial chains of vermis and hemisphere are aligned and the cortex between them usually is continuous. The cortex of the pyramis (Larsell's lobule VIII) 135

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continues into the caudal part of the paramedian lobule as the copula puramidis. The paraflocculus and the flocculus are the caudal segments of the folial chain of the hemisphere. The cortex between the paraflocculus and flocculus and the caudal vermis 136

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5mm. Fig. YY. Cerebellum of Mucucu fusciculuris. a. Anterior aspect. b. Ventral aspect. c. Caudal aspect. d . Dorsal aspect. Regions without cortex, where the white matter comes to the surface, are indicated with light hatching. Heavy hatching indicates cross section of the cerebellar peduncles, solid black indicates roof of the fourth ventricle in b. CrI = Crus I of the ansiform lobule; CrII = crus I1 of the ansiform lobule; FLO = flocculus; fpl = posterolateral fissure; PFLD = dorsal parafloculus; PFLV = ventral paraflocculus; PMD(cop) = paramedian lobule (copula pyramidis); S1 = lobulus simplex.

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J. Voogd, D. Jaarsma and E. Marani

(uvula and nodulus: lobules IX and X of Larsell) usually is completely interrupted. White matter invades the centre of the successive loops of the folial chain of the hemisphere and separates the successive segments of the flocculus and the paraflocculus. The dorsal and ventral segments of the paraflocculus are designated as the dorsal and ventral paraflocculus and the portion of the paraflocculus located in the subarcuate fossa of the petrosal bone is known as the petrosal lobule. This nomenclature is purely descriptive and these terms do not necessarily refer to identical segments of the paraflocculus in different species (see Fig. 97e for the topology in the rat and Fig. 149 for a comparison of rabbit, monkey and cat). The border between the paraflocculus and the flocculus is the posterolateral fissure. This fissure is one of the first to develop in the hemisphere. Its localization was established by Larsell (1970) for many mammalian species. Larsell tried to verify the relations between vermis and hemisphere by tracing the development of the transverse fissures. He attached great importance to the confluence of the vermal and hemispheral parts of fissures which, in the adult, would indicate the transverse continuity of the lobules of vermis and hemisphere. Larsell's nomenclature for the cerebellar hemisphere is rather unpractical and the significance of some of his many subdivisions (significance in the sense that a portion of the hemisphere has something in common with the vermal lobule to which it belongs) is questionable.

5. THE CEREBELLAR NUCLEI The (central) cerebellar nuclei and the lateral vestibular nucleus of Deiters receive the axons of the Purkinje cells of the cerebellar cortex and serve as the main output stations of the cerebellum. The vermis and the flocculus also project to other vestibular nuclei, but here the Purkinje cell axons compete with vestibular root fibers, intrinsic and commissural vestibular connections and projections from the medial cerebellar nucleus and, therefore, are not the dominant afferent system. Large numbers of Purkinje cells axons converge upon the cerebellar nuclei. The Purkinje cell/dentate nuclear cell convergence ratio is about 14:1 in the rhesus monkey and about 30:1 in the rat (Chan-Palay, 1977). It was estimated from Golgi impregnated Purkinje cell axons of the rat that between 20-50 perikarya of central nuclear neurons are included within the conical terminal field of a single axon (Chan-Palay, 1977), but the actual number of central nuclear neurons contacted by a single axon must be far greater. A considerable overlap in the corticonuclear projection of adjacent cortical areas was observed in axonal tracing studies (Courville and Diakew, 1976; Armstrong and Schild, 1978a,b; Haines and Koletar, 1979; Haines et al., 1982). Convergence and overlap are greatest in the rostrocaudal dimension where the entire length of the cortical sheet is compressed in the small volume of the cerebellar nuclei. Convergence is much less in the transverse direction and overlap may be even absent at the borders of neighbouring longitudinal zones where the Purkinje cells project to different nuclei (Voogd, 1964; Voogd and Bigar6, 1980). The convergence of the Purkinje cells from apex and base of a lobule onto the same cerebellar nuclear neurons is usually taken for granted. This type of convergence is of potential interest, because it combines the output of Purkinje cells that are under the influence of quite different mossy fiber-parallel fiber systems: corticopontine and exteroceptive systems at the apex and vestibular and proprioceptive systems at the base of the lobules (see also Section 6.4.2.). The subdivision of the cerebellar nuclei is closely related to the longitudinal, zonal 138

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organization of the corticonuclear and olivocerebellar projections. In this view the cerebellar nuclei are the output stations of corticonuclear modules and their independent olivocerebellar control systems. Certain cytochemical correlates of this longitudinal organization exist, but there are no signs of a corresponding cytochemical differentiation in the output of the cerebellar nuclei. The main input to the cerebellar nuclei from the Purkinje cells is GABAergic and inhibitory (Sections 3.1.1. and 5.5.). The nuclear cells receive their excitatory drive from collaterals from afferent mossy and climbing fiber systems (Section 5.6.). Collaterals of olivocerebellar fibers are organized according to the same longitudinal principle as their climbing fiber terminals, i.e. collaterals terminate in the cerebellar nucleus that receives axons from Purkinje cells that receive climbing fibers from the same parent axons (Groenewegen and Voogd, 1977). Mossy fiber input to the cerebellar nuclei is more diffuse, i.e. not limited to a single nucleus, and more selective, i.e. not present in all mossy fiber systems. Mossy fibers, therefore, are diverse in origin and dissimilar with respect to their collateral projections to the cerebellar nuclei, but as yet neurochemical correlates of this diversity are lacking. The monoaminergic input to the central nuclei 139

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J. Voogd, D. Jaarsma and E. Marani

is diffuse and independent of the mossy and climbing fiber collaterals. It is mainly represented by serotonin in a varicose, all pervading plexus (Section 5.7.). Neurons of the cerebellar nuclei are a mixed population of cells of all shapes and sizes. Several authors noticed a binominal distribution for cell size in the cerebellar nuclei. According to Courville and Cooper (1970) all four central nuclei of the monkey contain cells of all sizes. They noticed a peak in the size distribution for the lateral and interposed nuclei at 22 ~tm cell diameter. Palkovits et al. (1977) distinguished two populations of neurons, one with small and one with large nuclei, in the fastigial and posterior interposed nuclei and less clearly so in the lateral nucleus of the cat. Chan-Palay (1977) classified the cells of the lateral cerebellar nucleus in approximately equal groups of small and large neurons, with a separation at 180 t i m 2 surface area for the rat and 270 t i m 2 for the monkey. These binominal distributions can be explained by the presence of a population of small, GABAergic neurons and a non-GABAergic population of putative glutamatergic cells of different sizes, as recently reported by Batini et al. (1992) for the rat cerebellar nuclei (Section 5.1.3.) (Fig. 110). The targets of the projections of individual cerebellar nuclei in the brain stem and the cord differ, but some of their projections i.e. to the thalamus, show a remarkable degree of overlap. As yet there exists no corresponding differentiation in the chemoarchitecture of the cerebellar nuclei. The only system for which the neurotransmitter is known is the nucleo olivary projection that takes its origin from the population of small, GABAergic neurons (Mugnaini and Oertel, 1985). GABA also may be present in the intrinsic connections of the cerebellar nuclei and in their nucleocortical projection (Section 5.3.). 5.1. SUBDIVISION OF THE CEREBELLAR NUCLEI The cerebellar nuclei with their efferent tracts border on the ventricular surface of the cerebellum. Ventrolaterally they are continuous with the vestibular nuclei. They are surrounded by a sheet of afferent fibers from the restiform body and the middle cerebellar peduncle. This sheet is perforated by the bundles of Purkinje cell axons that terminate in the cerebellar nuclei or continue through or along these nuclei to the vestibular nuclei. The cerebellar nuclei of mammals can be subdivided according to Brunner (1919) into three, medio-laterally arranged nuclei or according to Weidenreich (1899) and Ogawa (1935) into a rostrolateral and a caudomedial nuclear group. Cytoarchitectonic criteria can be used to subdivide the cerebellar nuclei, but the presence of fiber bundles, and the disposition of their efferents in the nuclear hilus and in their efferent tracts are especially important in this respect. Brunner's (1919) mediolateral subdivision into the medial (fastigial), lateral (dentate) and interposed nuclei is based on the contours of the cerebellar nuclear mass. The three nuclei are part of a continuum and the nuclear borders therefore remain arbitrary. Although Brunner's concept of the central nuclei as a single mass has been disproved, his names for the nuclei have been retained. Weidenreich's (1899) comparative studies in mammals had already shown that fiber bundles subdivide the nuclei into a caudomedial group, that includes the medial nucleus and the caudal part of the Brunner's interposed nucleus (the nucleus interpositus posterior of Ogawa, 1935) and a rostrolateral group that is composed of the lateral cerebellar nucleus and the rostral part of the interposed nucleus (the nucleus interpositus anterior). The nuclei within each group are interconnected by cell bridges. The subdivision of Weidenreich-Ogawa can be readily appreciated in lagomorpha (Ono and Kato, 1938; Snider 1940) in carnivores (Flood and 140

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Jansen, 1961; Voogd 1964) and primates (Courville and Cooper, 1970; Chan-Palay, 1977). In rodents and insectivores the separation between the two nuclear groups is less distinct and the connections between the nuclei of each group are more extensive (Korneliussen, 1968a; Ohkawa 1957). The subdivision of Weidenreich-Ogawa received strong support from the localization of the fibers in the superior cerebellar peduncle. A small medial and a large lateral portion can be distinguished in this pathway in most mammals at its exit from the central nuclei (see* in Fig. 102). Experiments in cat (Verhaart, 1956; Voogd, 1964) and rat (Haroian et al., 1981) have shown that the medial part of the superior cerebellar peduncle takes its origin from the nuclei of the caudomedial group, mainly from the ipsilateral posterior interposed nucleus, and the lateral portion from the ipsilateral anterior interposed and lateral cerebellar nucleus. The efferent connections of the cerebellar nuclei were reviewed by Voogd et al. (1990), Ruigrok and Cella (1995), Voogd (1995) and Glickstein and Voogd (1995). The efferent connections of the lateral and interposed nuclei are summarized in Fig. 101. They give rise to nucleo olivary fibers which, at their exit from the nuclei, occupy a more ventral and medial position than the main bundle of the superior cerebellar peduncle (Legendre and Courville, 1987). After its decussation at the border of met- and mesencephalon, the superior cerebellar peduncle splits in ascending and descending branches. An uncrossed descending branch that detaches from the peduncle prior to its decussation, is only present in rodents (Ramon y Cajal, 1911). The main target of the crossed descending branch is the nucleus reticularis tegmenti pontis. This nucleus gives rise to a recurrent 141

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Fig. 102. A. Course of the direct fastigiobulbar tracts (left) and the uncinate tract (u) (right) in the stereotaxic planes P6-P8 in the cat. bc, brachium conjunctivum; bp, brachium pontis; CO, cochlear nuclei; cr, restiform body; dfb, direct fastigiobulbar tract; drV, spinal tract of the trigeminal nerve; DV, spinal vestibular nucleus; F, fastigial nucleus; FLO, flocculus; gVII, genu of the facial nerve; IA, anterior interposed nucleus; L, lateral cerebellar nucleus; LV, lateral vestibular nucleus; mlf, medial longitudinal fasciculus; MV, medial vestibular nucleus; sad, dorsal acoustic striae; SV, superior vestibular nucleus; vma, anterior medullaryvelum; vsc, ventral spinocerebellar tract. Modified from Voogd (1964). B. Hfiggqvist-stainedsection at the level of P7. Note the uncinate tract (u) arching over the brachium conjunctivum and the medial course of the direct fastigiobulbar tract (dfb). bc, brachium conjunctivum; cr, restiform body; dfb, direct fastigiobulbar tract; IA, anterior interposed nucleus; oc, olivocerebellar fibers; SV, superior vestibular nucleus; u, uncinate tract; vsc, ventral spinocerebellar tract; asterisk, medial one-third of the brachium conjunctivum, containing fibers from the posterior interposed nucleus. Voogd et al. (1990) (

mossy fiber pathway to the cerebellar cortex, with a strong collateral projection to the cerebellar nuclei. The ascending branch terminates in the red nucleus, in nuclei at the mesodiencephalic junction, including Darkschewitsch nucleus, that give rise to the medial and central tegmental tracts, which terminate in the inferior olive. The ascending branch terminates in the thalamus. Not all the nuclei contribute equally to each connection. The lateral and the anterior interposed nuclei project heavily to the nucleus reticularis tegmenti pontis and the red nucleus, whereas the posterior interposed nucleus is preferentially connected with the nuclei of the mesodiencephalic junction. The fastigial nucleus gives rise to fibers of the uncinate tract, that decussate within the cerebellum, and to the ipsilateral fastigiobulbar tract, that passes medial to the superior cerebellar peduncle in the lateral wall of the fourth ventricle (Fig. 102) (Voogd, 1964; Batton et al., 1977). The uncinate tract gives rise to a small, ascending bundle, that terminates in the central grey, the mesencephalic tegmentum and the thalamus. The majority of the uncinate- and direct fastigiobulbar tract fibers terminate in the vestibular nuclei and the reticular formation of pons and medulla oblongata. Regions containing predominantly small cells have been distinguished in the ventral parts of the cerebellar nuclei of cat (Flood and Jansen, 1961; Voogd, 1964), rat (Korneliussen, 1968a; Beitz and Chan-Palay, 1979; Voogd, 1995), several subprimates (Haines, 1977b) and primates (Courville and Cooper, 1970). These parvicellular regions are not well-defined and difficult to compare in different species. Even where they were designated as subnuclei (Flood and Jansen, 1961) these subdivisions had no clear functional basis. Data on the connections of the small cells are scanty and conflicting. Haines (1977b) proposed direct and indirect projections of the parvicellular medial and lateral cerebellar nuclei to the oculomotor nuclei. Itoh and Mizuno (1979) found a projection of the parvicellular dentate in the cat to the pulvinar and Mugnaini and Oertel (1985) noticed a concentration of small, GABAergic, nucleoolivary neurons in the ventral parvicellular part of the lateral nucleus of the rat. The group y of Brodal and Pompeiano (1957) and the basal interstitial nucleus of Langer (1985) are two nuclei which are located within the cerebellum, outside the 'classical' cerebellar nuclei. The group y was defined as a small-celled subgroup of the vestibular nuclei of the cat, that caps the restiform body dorsally. Ventrolaterally it is in contact with the dorsal cochlear nucleus. Dorsally scattered cells form strands extending to the dentate nucleus. Medially it is continuous with the small cells of the caudal pole of the superior vestibular nucleus, located dorsolateral to Deiters' nucleus also known as the group 1 of Brodal and Pompeiano (1957). In transverse sections the cells of the y group are fusiform because they are located between the fibers of the floccular peduncle (Voogd, 1964). 143

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144

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Fig. 104. Two transverse, AChE-incubated sections through the cerebellar nuclei of the cat. A. Rostral section. B. Caudal section. Note medium-sized cells of dorsal group y in floccular peduncle and strongly AChE positive ventral group y along dorsal border of restiform body in (A); U-shaped nucleus between IP and F in (B). cr = restiform body; F = fastigial nucleus; flo+y = floccular peduncle with group y; IA = anterior interposed nucleus; IP - posterior interposed nucleus; IP/F = U-shaped nucleus between F and IP; L = lateral cerebellar nucleus; sad = stria acoustica dorsalis.

Graybiel and Hartweg (1974) found retrograde labelling in the cells of the y group of the cat after injection of retrograde tracers in the oculomotor nucleus. Matters have been further complicated by the introduction of the term 'infracerebellar nucleus' for the y group by Gacek (1977, 1979)l, and the distinction of ventral and dorsal divisions in the group y (Kevetter and Perachio, 1986; Highstein and Reisine, 1979; Highstein and 145

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J. Voogd, D. Jaarsma and E. Marani

Fig. 105. The cerebellar nuclei of Macaca fascicularis. Upper diagram is a graphical reconstruction of the cerebellar nuclei in a dorsal view. Levels of the transverse sections are indicated. The U-shaped, transitional nucleus located between the fastigial (F) and posterior inter-posed nucleus (IP) is indicated with double hatching, bc = brachium conjunctivum; BIN = basal interstitial nucleus of Langer; cr = restiform body; DV = descending vestibular nucleus; F = fastigial nucleus; IA - anterior interposed nucleus; IP = posterior interposed nucleus; L = lateral cerebellar nucleus; LV = lateral vestibular nucleus (Deiters'); MV = medial vestibular nuleus; SV = superior vestibular nucleus; Y = group y; asterisk = medial one-third of the brachium conjunctivum. )

McCrea, 1988). A more realistic approach would be to consider the cells of the y group as the bed-nucleus of the floccular peduncle, and to realize that a distinction between the group y and the superior vestibular nucleus, that receives a major part of the fibers of the floccular peduncle, remains arbitrary (Tan et al., 1995a). The basal interstitial nucleus was described by Langer (1985) in the monkey as a broadly distributed interstitial population of neurons in the white matter ventral to the cerebellar nuclei and extending from the white matter of the nodulus, into the peduncle of the flocculus. The nucleus was distinguished from the group y in the monkey. It is reciprocally connected with the flocculus. It seems likely from their descriptions that the group y of Brodal and Pompeiano corresponds to the portion of Langer's basal interstitial nucleus that lies embedded in the floccular peduncle. The 'group y' of the monkey can be considered as the enlarged, caudal pole of the superior vestibular nucleus (corresponding to the group 1 of Brodal and Pompeiano, 1957, in the cat). Some features of the cerebellar nuclei of the cat, the monkey, and the rat shall be discussed in the next sections. The cerebellar nuclei of birds were described by Feirabend (1983) and Arends and Zeigler (1991 a,b). 5.1.1. The cerebellar nuclei of the cat (Figs 103 and 104) The subdivision of Weidenreich-Ogawa can be applied to the central nuclei of the cat. This is not amazing, because Ogawa's (1935) description of the central nuclei of pinnepedia is based on earlier, unpublished, material from the cerebellum of cat and dog. Moreover the parasagittal organization of the corticonuclear and olivocerebellar projections, that provided important clues for the subdivision of the central nuclei, was first and most extensively studied in the cat (Hohman, 1929; Voogd, 1964, 1969; Courville et al., 1974; Groenewegen and Voogd, 1977). Large cells are prominent in the rostral part of the medial nucleus, small cells predominate in its ventromedial and caudal parts (Flood and Jansen, 1961). The lateral border of the medial nucleus is flush with the AChE-positive raphe which forms the lateral border of the medial A-compartment of the anterior vermis. AChE is concentrated in the lateral and ventral parts of the medial nucleus and in the neuropil of cell groups scattered in between the medial and the anterior interposed nucleus. Caudally these AChE-positive clusters coalesce in a U-shaped nucleus located at the transition of the medial and the posterior interposed nucleus (Fig. 104B). The medial limb of the U forms the lateral border zone of the fastigial nucleus, the lateral limb usually is included with

1 Gacek (1977, 1979) used the term infracerebellar nucleus for the cells embedded in the fasciculus angularis (i.e. the floccular peduncle) as it arches over the restiform body and applied the term 'y group' to the cells of the group 1 of Brodal and Pompeiano (1957).

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the posterior interposed nucleus. The large cells in this area are the target of the Purkinje cells in the X-zone of the anterior vermis (Trott and Armstrong, 1987a,b). The cells in the lateral limb of the U, that extend rostrally as the AChE-positive cell groups medial to the anterior interposed nucleus, may provide the target for the 'C ~' zone, the lateral portion of the Cl-zone, that shares its afferent, climbing fiber input with the X-zone (Ekerot and Larson, 1982; Campbell and Armstrong, 1985; Trott and Armstrong, 1987a) (Section 6.3.3.1., Fig. 174). The cells of this area project to the spinal cord (Matsushita and Hosoya, 1979) and can be double-labelled from the cord and the thalamus (Bharos et al., 1981). Apart from the medial limb of the U-shaped subnucleus, the medial nucleus at this level consists of a dorsally directed tail that extends into the white matter of the posterior lobe vermis. It receives Purkinje cell axons from the vermal visual area of lobule VII (Voogd, 1964; Courville and Diakew, 1976). It constitutes one of the targets of the collateral projection from the nucleus reticularis tegmenti pontis (Gerrits and Voogd, 1987). Other parts of the medial nucleus receive their mossy fiber collaterals from the lateral reticular nucleus (Ktinzle, 1975; Russchen et al., 1976) and the spinal cord (Matsushita and Ikeda, 1970). The ventral, parvicellular part of the medial nucleus is located at the base of the lobules IX and X. The posterior and anterior interposed and the lateral cerebellar nucleus are clearly delimited. The border between the anterior and posterior interposed nucleus is not located in a frontal plane, but passes obliquely forward, from caudolaterally to rostromedially (Fig. 103). The posterior interposed nucleus, therefore, extends far rostrally, medial to the anterior interposed nucleus. The caudal pole of the anterior interposed nucleus is located far laterally, adjacent to and merging with the lateral nucleus. AChE is concentrated along the borders of the nuclei and in the ventrolateral portions of the lateral and posterior interposed nuclei. Elongated, AChE-positive cells in the floccular peduncle, ventral to the lateral nucleus, belong to cell group y of the vestibular nuclei of Brodal and Pompeiano (1957) (Fig. 104A). The medium-sized cells of group 1 which are located dorsolateral to Deiters' nucleus, should be distinguished from the group y. They constitute a caudal extension of the superior vestibular nucleus and display strong AChE activity.

5.1.2. The eerebellar nuclei of primates The cerebellar nuclei in primates were described by Courville and Cooper (1970) and Chan-Palay (1977, Macaca mulatta), Haines (1971, Galago), and Haines and Dietrichs (1991, Saimiri sciurus). The cerebellar nuclei of the human cerebellum were reviewed by Larsell and Jansen (1972) and Voogd et al. (1990). The four nuclei of the subdivision of Weidenreich-Ogawa can be recognized in macaque fascicularis and their topograph-

Fig. 106. Horizontal (A) and transverse (B,C) AChE-stained sections through the cerebellar nuclei of Macaca fascicularis. Note connection of IA and dorsal pole of dentate nucleus in (B) and of medial lamella of the dentate nucleus and border region of IA and IP in (A), U-shaped nucleus located between F and IP in (B) and (C) and strong AChE-reactivity in this nucleus in (A); localization of medial limb of this U-shaped nucleus in X compartment (B and C); extension of AChE-positive C2 compartment in border region of 1A and IP in B; interstitial nucleus of Langer in (B) and large group y in (C). C2 = C2 compartment; F = fastigial nucleus; IA = anterior interposed nucleus; IP = posterior interposed nucleus; IP/F = U-shaped nucleus between F and IP; L = lateral cerebellar nucleus; Lp = parvocellular part of lateral cerebellar nucleus; y = group y; X = X compartment. 148

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of a core and a shell; the neuropil of the shell is connected with the medial limb of the dentate nucleus. The anterior interposed nucleus consists of medial and lateral portions, separated by a notch in the dorsal surface. The caudal pole of the anterior interposed nucleus merges with the medial limb of the dentate nucleus. The dentate nucleus displays ventrolaterally and caudally directed bulges. The hilus of the dentate nucleus is directed rostromedially, caudally it is closed by the medial limb of the dentate nucleus. In its caudal portion the structure of the medial limb is compact, with AChE-positive cell bodies and neuropil concentrated along its borders, similar to the rest of the dentate nucleus. Rostrally where the medial limb surrounds and merges with the anterior interposed nucleus, its structure is loose and AChE-staining less prominent, A subdivision of the dentate nucleus of the monkey in the two parts that were distinguished in the human dentate nucleus, i.e. in the rostromedial neodentatum with large neurons and the caudolateral palaeodentatum, with densely packed, smaller neurons (Gans, 1924; Demol6, 1927a,b) is not feasible and is not warranted by Chan-Palay's (1977) detailed cytological analysis of this nucleus. The so-called group y is large and compact and occupies the rostral part of the floccular peduncle. Medially it is continuous with the caudal pole of the superior vestibular nucleus. The small AChE-positive cells of Langer's (1985) basal interstitial nucleus appear more caudally and extend from the flocculus into the roof of the fourth ventricle, ventral to and in between the dentate and posterior interposed nuclei. 5.1.3. The cerebellar nuclei of the rat

The cerebellar nuclei of the rat were described by Goodman et al. (1963), Korneliussen (1968a) and Voogd (1995). They are very similar to the nuclei of the cerebellum of the mouse, illustrated by Marani (1982a). The morphology and the cytology of the lateral and the medial nucleus of the rat were analysed in Chan-Palay's (1977) monograph on the dentate nucleus and in the paper of Beitz and Chan-Palay (1979). The cerebellar nuclei of the rat are difficult to subdivide according to the scheme of WeidenreichOgawa. They constitute, more or less, a single mass, with a number of protrusions, that were first named by Goodman et al. (1963) as the dorsolateral protuberance of the medial nucleus, and the dorsomedial crest and the dorsolateral hump, two excrescences of the interposito-dentate complex (Figs 107 and 108). Similar subnuclei were distinguished in the cerebellum of the mouse (Marani, 1986). The connections of the cerebellar nuclei in the rat differ in some respects from carnivores and primates. The medial nucleus was subdivided by Korneliussen (1968a) into a caudoventral parvicellular part, the magnocellular dorsolateral protuberance and a 'middle' part that contains cells of intermediate size. The dorsolateral protuberance is situated more laterally. It projects into the white matter of the posterior lobe and receives Purkinje cell fibers from a medial zone of the lobules VI and VII of the posterior lobe hemisphere (Goodman et al., 1963; Armstrong and Schild, 1978a,b; Buisseret-Delmas, 1988a,b) (see also Section 6.1.4.). The posterior interposed nucleus is best distinguished in horizontal sections. A cellfree zone separates it from the anterior interposed and lateral nuclei. Medially the posterior interposed nucleus is continuous with the fastigial nucleus. Cells at the junction of these two nuclei project to the spinal cord (Matsushita and Hosoya, 1978 and Bentivoglio and Kuypers, 1982). This region was distinguished as the interstitial cell group, the target nucleus of the X zone, by Buisseret-Delmas et al. (1993). At the junction of the anterior interposed and lateral nuclei the dorsolateral hump 151

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Fig 108. The cerebellar nuclei of the rat. Left, Nissl stain; right, AChE-incubations of adjacenl transverse sections. Note border between scattered cells of rostra1 pole of F and AChE-positive region in the lateral bank of the fourth ventricle in (B); strong AChE staining in caudal pole SV, group y and parvocellular dentate in (D); border between strongly AChE positive D L H and medium staining of IP in (F) and cell strands of caudal pole of F in two AChE-positive raphes (F). bc = brachium conjunctivum; bp = brachium pontis; C O = cochlear nuclci; cr = restiform body ; D L H = dorsolateral hump; D L P = dorsolateral protuberance of the fastigial nucleus; F = fastigial nucleus; flo+y = floccular peduncle with group y; 1A = anterior interposed nucleus; IP = posterior interposed nucleus; L = lateral cerebellar nucleus; Lp = parvocellular part of lateral cerebellar nucleus; LV = lateral vestibular nucleus of Deiters; MVm = magnocellular part of medial vestibular nucleus; MVp = parvocellular part of medial vestibular bucleus; SV = superior vestibular nucleus; V = spinal tract of trigeminal nerve; XI11 = vestibular nerve; * = medial one-third of the brachium conjunctivum.

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forms a distinct ridge on the rostral aspect of these nuclei. A more caudally located cluster of large neurons usually is indicated by the same name. Some authors included the dorsolateral hump with the anterior interposed nucleus (Goodman et al. 1963; Voogd, 1995), others delegated it to the lateral nucleus (Chan-Palay, 1977; Angaut and Cicirata, 1982), sometimes it was considered as a separate subdivision, peculiar to rodents. It gives rise to the uncrossed descending limb of the superior cerebellar peduncle (Woodson and Angaut, 1984; Bentivoglio and Molinari, 1986; Rubertone et al., 1990), that detaches from the superior cerebellar peduncle before its decussation. The uncrossed descending limb terminates in the lateral reticular formation (Ramon y Cajal, 1911; Mehler, 1967, 1969; Achenbach and Goodman, 1968; Faull, 1978; Chan-Palay, 1977). The ventromedial part of the lateral nucleus consists of smaller cells. Those in the region of the hilus correspond to the small intrinsic neurons described by Chan-Palay (1977), the densely packed small cells located ventral to the hilus may be identical to the concentration of GABAergic nucleo-olivary cells that was illustrated in this region by Mugnaini and Oertel (1985). AChE staining does not distinguish the parvicellular part of the lateral nucleus, but the medium-sized cells and the neuropil of the group y, located ventral to it display a high AChE activity. In a rostral direction the group y (Brodal and Pompeiano, 1957) becomes continuous with the superior vestibular nucleus). Deiters' nucleus with its large AChE-positive perikarya in an unstained neuropil is wedged in between the AChE-rich areas of the group y and the medial vestibular nucleus and reaches far dorsally into the hilus region of the central nuclei. Purkinje cell fibers enter Deiters' nucleus as perforating fibers, passing in between the dorsolateral protuberance and the anterior interposed nucleus, and through the middle part of the medial nucleus. More rostrally, where Deiters' nucleus has disappeared, the AChE-rich neuropil of the superior and medial vestibular nuclei meet at the oblique border between the two nuclei. 5.2. THE GABAERGIC NUCLEO-OLIVARY PROJECTION NEURONS OF THE CEREBELLAR NUCLEI Cell size has generally been considered of less importance for the subdivision of the central nuclei, than the presence of local differences in cell density resulting from sheets or bundles of fibers between and around the nuclei. Interest in the question whether different cell-types can be distinguished in the cerebellar nuclei on the basis of size, dendritic and axonal morphology and neurotransmitter content was renewed since the observation that the projection of the cerebellar nuclei to the inferior olive arises from a population of small GABAergic neurons. The nucleo-olivary projection was discovered with anterograde tracing with tritiated amino acids by Graybiel et al. (1973) and with retrograde tracing by Gould and Graybiel (1976) and Tolbert et al. (1976b), all in the cat. The nucleo-olivary and olivo nuclear projections are reciprocally organized (see Section 6.3.3.). Tolbert's (Tolbert et al., 1978a) and Courville and Cooper's (1970) quantitative analysis clearly showed that all sizes of neurons were present in all central nuclei in monkey and cat. Histograms of the soma diameter of the nuclear neurons projecting to the thalamus and the cerebellar cortex are very similar to the overall size distribution of these neurons. The cells in the cerebellar nuclei of the cat that project to the inferior olive, however, constitute a population of small, spindle shaped neurons (Fig. 109). This 154

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population of small cells was recognized in retrograde tracer experiments in opossum (Martin et al., 1976, 1980) cat (Tolbert et al., 1976b; Tolbert et al., 1978a; Bharos et al., 1981; Legendre and Courville, 1987), rat (Angaut and Cicirata, 1982; Hess, 1982b; Brown et al., 1977; Bentivoglio and Kuypers, 1982; Buisseret et al., 1989; Swenson and Castro, 1983b) and monkey (Chan-Palay, 1977). The small cells were depleted in the cerebellar nuclei of lurcher mutant mice, probably due to the degeneration of the inferior olive (Heckroth, 1994). According to Tolbert et al. (1978a) the nucleo-olivary neurons in the cat are concentrated in the ventral parts of the dentate and posterior interposed nuclei, they are scarce in the fastigial nucleus. Concentrations of these small neurons were also reported in the rostral and caudal poles and the hilar portion of the dentate nucleus and the lateral parts of the interposed nucleus in the rat (Brown et al., 1977; Chan-Palay, 1977). A more diffuse distribution of these cells was noticed in experiments with retrograde tracing of Martin et al. (1976) in the dentate and interposed nuclei of the opossum and in double labelling studies with fluorescent dyes in rat (Bentivoglio and Kuypers, 1982) and cat (Bharos et al., 1981). Neurons of the cerebellar nuclei and the lateral cerebellar nucleus of rat and cat that react with antibodies against GAD or conjugates of GABA, generally are small (Mugnaini and OerteI, 1981, 1985; Houser et al., 1984; Gabbott et al., 1986; Kumoi et al., 155

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Fig. 110. Diameter distributions of GABA-like immunoreactive (GABA-IR) and glutamate-like immunoreactive (Glu-IR) neurons in the nucleus medialis (NM), the nucleus interpositus (NI) and the nucleus lateralis (NL) of rat cerebellum. The populations of GABA-IR (A) and Glu-IR (B) neurons were clearly peaked and similar in size in all three nuclei. In C the spectra from the three nuclei are averaged for both populations and plotted together. The size range of the GABA and Glu overlap is the same as for cells positively identified as colocalizing GABA and Glu. Abscissae: diameter of the neurons in p m (class interval 2.5/lm). Ordinates: percentage of neurons in each diameter class. Batini et al. (1992).

1987, 1988; Buisseret-Delmas et al., 1989; Walberg et al., 1990; Takayama, 1994; Moffett et al., 1994). Batini et al. (1992), who measured cell diameters of GABA and single labelled glutamate-immunoreactive neurons, showed that they represent two populations with little overlap in all cerebellar nuclei of the rat (Figs 110 and 111). Consistent with glutamate being a metabolic precursor for GABA, most of the GABAergic neurons co-localized glutamate. GAD or GABA-like immunoreactive cells are scattered through all parts of the cerebellar nuclei, but have been reported to be more sparse in the fastigial nucleus, where they are concentrated in its ventral portion and to be preferentially located in the hilus and in the ventral parvocellular subnucleus of the lateral cerebellar nucleus of the rat (Mugnaini and Oertel 1985; Buisseret-Delmas et al. 1989). These small 156

The cerebellum." chemoarchitecture and anatomy

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Fig. 111. Typical distribution of glutamate-like and GABA-like immunoreactive neurons in the nucleus lateralis of rat cerebellum. Consecutive sections immunostained for glutamate (A) and GABA (B) are shown as mirror images. Note the difference in size of the stained neurons. Batini et al. (1992).

neurons proved to be the origin of the GABAergic, nucleo-olivary pathway in the rat (Nelson et al. 1984) and the rabbit (Nelson and Mugnaini, 1989). De Zeeuw et al. (1988, 1989b) demonstrated that all the terminals in the inferior olive that could be traced from the cerebellar nuclei by anterograde transport of WGA-HRP could be labelled with a polyclonal antibody against GABA. The nucleo-olivary projection, therefore, seems to be entirely GABAergic. Some of the small GABAergic perikarya of the cerebellar nuclei and the lateral vestibular nucleus also contain glycine (Ottersen et al., 1987; Walberg et al., 1990; Chen and Hillman, 1993b; Takayama, 1994), but glycine could not be detected in terminals of the nucleo-olivary pathway in the rat (De Zeeuw, unpublished observations). Projections to certain subdivisions of the inferior olive also take their origin from the vestibular nuclei (Saint Cyr and Courville, 1979; Gerrits et al. 1985a) that contains a population of small GABAergic neurons in all its subnuclei (Kumoi et al., 1987, guinea pig). Some of these connections are also GABAergic (Nelson et al., 1986), but it is not known whether all vestibulo-olivary connections use this neurotransmitter. One of the main sources for the vestibulo-olivary projection is the nucleus prepositus hypoglossi (Saint Cyr and Courville, 1979; McCrea and Baker, 1985; Gerrits et al., 1985a). Both GABA (de Zeeuw et al., 1993) and acetylcholine (Barmack et al., 1991) have been identified as neurotransmitters in this pathway. At present it seems unlikely that the small nucleo-olivary neurons possess collaterals that terminate in other targets. Tolbert et al. (1978a), however, found many of these neurons in the cat to be antidromically activated from the thalamus and the cerebellar cortex in addition to the inferior olive. Ban and Ohno (1977) also produced electrophysiological evidence for collateralization of nucleo-olivary cells to the red nucleus or more rostral structures. Bharos et al. (1981) and Bentivoglio and Kuypers (1982), were unable 157

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to double label these small neurons, after combined injections of one fluorescent tracer in the caudal medulla, in a region including the inferior olive, and a second injection with another fluorescent tracer in either thalamus with tectum or the spinal cord. These results were corroborated for the combination of thalamus and inferior olive by Legendre and Courville (1987) for the cat and the red nucleus and the inferior olive by Teune et al. (1995) for the rat. The existence of GABAergic projections of the cerebellar nuclei to other precerebellar relay nuclei is not excluded by these experiments. Projections from GABAergic neurons of the lateral cerebellar nucleus of the rat to the basal pontine nuclei were demonstrated by Border et al. (1986). 5.3. NUCLEOCORTICAL AND INTRINSIC NEURONS OF THE CEREBELLAR NUCLEI Nucleocortical neurons occasionally have been described in Golgi material (Ramon y Cajal, 1911) and were considered as the origin of the climbing fibers by Carrea et al. (1947). Their existence was confirmed by retrograde transport of HRP injected in various parts of the cerebellar cortex in the cat (Gould and Graybiel, 1976; Tolbert et al., 1976a, 1978b; Dietrichs, 1981a,b, 1983a; Dietrichs and Walberg, 1979a, 1980; Trott and Armstrong, 1990), in primates (Tolbert et al., 1977, 1978b; Tolbert and Bantli, 1979; Haines, 1978b, 1988; Haines and Pearson, 1979), and in the rat (Chan-Palay, 1977; Hess, 1982a; Buisseret-Delmas and Angaut, 1988, 1989a). The nucleocortical projection was reviewed by Tolbert (1982). The size of the nucleocortical cells in the cat follows the same frequency distribution as the central nuclear cells as a whole and as the neurons that could be retrogradely labelled from the thalamus (Fig. 109). Single neurons, intracellularly stained with HRP, with axons leaving the central nuclei in the superior cerebellar peduncle, were shown to emit collaterals, that could be traced to the cerebellar cortex (McCrea et al., 1978). Antidromic invasion and collision experiments (Tolbert et al., 1977, 1978a) also favoured a collateral origin of the nucleocortical fibers from projection neurons. There is some evidence for GABA or acetylcholine as the neurotransmitter of the nucleocortical fibers, but the majority, probably, uses glutamate or aspartate, in accordance with their collateral origin from large relay cells of the cerebellar nuclei. 20% of the nucleocortical neurons in the medial cerebellar nucleus that were WGA-HRP labelled from the cerebellar cortex of the rat, reacted with an antibody raised against GABA (Angaut et al., 1988). Most if not all nucleocortical neurons in the rat were found to be immunoreactive for an antibody to a conjugate of GABA in the experiments of Batini et al. (1989). She labelled GABAergic cells of all sizes from the cortex, although fewer retrogradely labelled cells were present among the smaller GABAergic neurons, most of which presumably project to the inferior olive. The proportion and the size distribution of nucleocortical neurons of the rat labelled with the same antibody to a conjugate of GABA were reported to be different in a later publication of the same authors (Batini et al., 1992). Again, the retrogradely labelled nucleocortical neurons were of all sizes with a peak around a diameter of 20-25 r With almost 50%, the population of single labelled glutamate-immunoreactive nucleocortical neurons far exceeded the GABAergic population. The glutamate-immunoreactive neurons were also larger and peaked at a diameter of 20-25 r The GABAergic-nucleocortical cells may be identical to the small nucleocortical neurons that were identified in the posterior interposed nucleus and along the boundaries of the lateral nucleus of the rat by retrograde transport of an antibody to GAD 158

The cerebellum." chemoarchitecture and anatomy

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(Chan-Palay et al., 1979). Antegrade transport of an antibody to GAD in nucleocortical fibers resulted in labelling of a small number of mossy fiber rosettes in the cortex (Chan Palay et al., 1979). Antegrade transport in nucleocortical fibers of other, non-specific tracers also resulted in the labelling of mossy fiber rosettes (Kultas-Illinsky et al., 1979; Tolbert et al., 1980). The morphology of these nucleocortical mossy fiber rosettes was not distinctive and they never displayed any of the features of GABAergic terminals, such as pleomorphic vesicles and symmetrical synapses. H~mori and Tak~cs (1989) and H~.mori et al. (1990) distinguished 4 types of mossy fiber rosettes in the cerebellar cortex of rat and cat with immunohistochemistry of glutamate and GABA and deafferentation of the cerebellum. Two of their types of mossy fiber rosettes, i.e. large, GABA-immunoreactive rosettes, with round or pleomorphic synaptic vesicles that accounted for 3% of all mossy fiber rosettes in the cat and small rosettes with small, pleomorphic synaptic vesicles, that reacted with an antibody to a conjugate of glutamate and accounted for less than 10% of the mossy fiber rosettes in the rat, presumably represented the endings of nucleocortical fibers. Mossy fibers of extracerebellar origin terminated as large, glutamate-immunoreactive rosettes; a fourth type of small, GABAergic rosettes with small synaptic vesicles was of intracortical origin. According to Ikeda et al. (1991) the cholinergic, ChAT-immunoreactive afferent fibers in the cortex of the cerebellum of the cat, that include mossy fibers, disappear after electrolytic or kainate lesions of the cerebellar nuclei. Different types of nucleocortical neurons have been distinguished on the basis of their localization with respect to the corticonuclear projection. Most nucleocortical cells were considered to be of a reciprocal type since they were located within an area receiving Purkinje cell projections from the same region of the cortex from which they can be retrogradely labelled (Dietrichs, 1981a,b, 1983a; Dietrichs and Walberg, 1979a, 1980; Tolbert, 1982; Buisseret-Delmas and Angaut, 1988, 1989a; Haines, 1988). Other nucleocortical neurons were found outside the corticonuclear projection area, either ipsilaterally (the non-reciprocal neurons of Buisseret-Delmas and Angaut, 1988, 1989a, see also Hess, 1982a, Dietrichs and Walberg, 1979a, 1980) or even contralaterally (symmetrical neurons: Buisseret-Delmas and Angaut, 1988, 1989a rat; Tolbert et al., 1978b cat and monkey; Haines, 1978a,b; Haines and Pearson, 1979, treeshrew; Haines, 1988, Galago; Dietrichs, 1983a; Dietrichs and Walberg, 1980, cat). Non-reciprocal and contralateral neurons in cat and rat were most numerous in the fastigial and posterior interposed nuclei. Contralateral neurons in the posterior interposed nucleus of Galago were small and fusiform and resembled the nucleo-olivary neurons (Haines, 1988). Non-reciprocal connections were the rule in primates where they originated mainly from the ventral part of the dentate nucleus. Reciprocity in general was more noticeable in cat and rat. A more restricted origin of the sparse nucleocortical projection to the electrophysiologically identified C~ and C2 zones and the strong bilateral projection to the C2 zone of the paravermal cortex of the cat from the posterior interposed and fastigial nuclei was advocated by Trott and Armstrong (1990). The existence of a rest group of neurons that remains unaffected by large lesions of the efferent cerebellar pathways in the kitten has been claimed as evidence in favour of the presence of intrinsic or nucleocortical neurons in the central nuclei (Jansen and Jansen 1955). Many of these neurons were found to be large and to be located in the posterior interposed nucleus. Intrinsic neurons of the cerebellar nuclei have been observed in Golgi preparations of the rat by Chan-Palay (1973a, 1977) as small multipolar neurons in the dentate nucleus. The terminals of these intrinsic, inhibitory neurons on the soma and dendrites of cerebellar nuclear cells were tentatively identified as small 159

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boutons with few elliptical vesicles in a light matrix (E-type, Chan-Palay, 1973b, 1977). Wassef et al. (1986), however, were unable to distinguish between these E-types and Purkinje cell terminals (Chan-Palay's A-type) in the cerebellar nuclei of mutant Pcd mice. Some 15% of the small and large GAD-positive terminals in the cerebellar nuclei of these mutants were spared, although these mice lost 99% of their Purkinje cells. Intrinsic GABAergic connections of the nuclei should have been spared in these animals, but the possibility that these 15% represent sprouting from another GABAergic cell population cannot be excluded. Non-GABAergic intrinsic connections through boutons with spherical vesicles carrying Gray-type I synapses were postulated by Chan-Palay (D-type, 1973b); no other evidence is available on this system. Strong evidence for the presence of a population of small glycinergic interneurons in the cerebellar nuclei of the rat was supplied by Chen and Hillman (1993b). Glycine immunoreactivity was restricted to a population of small neurons throughout the cerebellar nuclei and was present in boutons outlining somata of large relay cells. Intense glycine receptor immunoreactivity was observed in these large cells, opposite the glycinergic terminals. Many, but not all of these small, glycinergic neurons colocalize GABA. Since glycine-immunoreactivity was not observed in nucleo-olivary terminals (De Zeeuw, unpublished observations) it seems likely that these glycine and glycine/GABA containing neurons are interneurons. Rather similar observations were made for calretinin in a subpopulation of small nuclear cells, that give rise to a dense plexus in the cerebellar nuclei of the rat (Floris et al., 1994). 5.4. NON-GABAergic PROJECTION N E U R O N S OF THE CEREBELLAR NUCLEI

Glutamate and aspartate have been suggested as the neurotransmitter of the nonGABAergic projection neurons. Immunoreactivity with an antibody to a conjugate of glutamate was found in neurons of all sizes in all cerebellar nuclei of the rat (Batini et al., 1992) (Fig. 110) and with antibodies to conjugates of aspartate in large neurons of the cerebellar nuclei of the rat (Kumoi et al., 1988; Chen and Hillman, 1993b). A reaction product of glutamate with carbodiimide (gamma-Glu-Glu) has been localized with immunocytochemistry in neurons of all sizes in all the cerebellar nuclei of carbodiimide-perfused rats (Monaghan et al., 1986, see also Madl et al., 1986, 1987). Moderate, glutaminase (GLNase)-like immunoreactivity was present in scattered small cells of the cerebellar nuclei of the rat (Kaneko et al., 1989). Cells reacting with an antibody against aspartate aminotransferase (AATase), that catalyses the conversion of glutamate to aspartate, are more numerous than the GLNase and gamma-Glu-Glucontaining cells, but not all cerebellar nuclear neurons were found to contain AATase (Monaghan et al., 1986). Since AATase-like immunoreactivity has been found in Purkinje, basket and stellate cells of the cerebellar cortex, this enzyme apparently is also present in GABAergic neurons. Monaghan et al. (1986) tentatively concluded that three types of neurons can be distinguished in the cerebellar nuclei of the rat. The first group includes putative glutamatergic neurons containing GLNase- and gamma-Glu-Glu-like immunoreactivity, the second group corresponds to the GABAergic neurons that contain AATase-like activity and the third group are the neurons that do not react with any of the three antibodies and whose neurotransmitters remain to be determined. They could not exclude the presence of aspartate-containing neurons in the cerebellar nuclei because elevated levels of AATase and GLNase can be present both in aspartatergic as well as in glutamatergic neurons. The large cells of Deiters' nucleus in guinea pig (Kumoi 160

The cerebellum." chemoarchitecture and anatomy

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Fig. 112. Retrograde transport of [3H]choline (A,B) to perikarya (arrow-heads) in medial part of nucleus interpositus (Int) or in lateral cerebellar nucleus (LatC) and of wheat germ agglutinin-coupled HRP (E,F), but absence of retrogradely labelled elements in corresponding regions after application of D-[3H]aspartate to the red nucleus (C,D). Rat, bright-field (A,E) as well as dark-field (B,F) illumination. Cresyl violet counterstaining. Bars = 0.5 mm. Bernays et al. (1988).

et al., 1987) and cat (Walberg et al., 1990), reacted with antibodies raised against conjugates of aspartate. The same giant cells in the cat also stained with antibodies to conjugates of glutamate (Walberg et al., 1990). The presence of excitatory amino acid transmitters in the projection neurons of the cerebellar nuclei was also investigated by retrograde transport of D-[3H]aspartate in the cerebellorubral pathway, but no such transport was observed in the rat (Bernays et al., 1988 (Fig. 112C,D)). Interruption of this pathway, however, resulted in a decrease of high affinity glutamate uptake in the caudal, magnocellular portion of the red nucleus (Nieoullon et al., 1984), which could be explained by a loss of cerebellorubral, glutamate containing terminals. High affinity uptake of glutamate in other cerebellar target nuclei 161

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The cerebellum." chemoarchitecture and anatomy

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Fig. 113. A. Darkfield micrograph of a frontal section through the cerebellar white matter and nuclei of an experiment with an injection of Phaseolus vulgaris leucaglutinin into part of the principal nucleus of the inferior olive and the rostral dorsal accessory olive in the rat. Labelled axons can be seen in the restiform body (cr). Bundles of labelled axons directed toward the cerebellar hemispheres seem continuous with innervated areas of the lateral cerebellar nucleus (L), the dorsolateral hump (DLH) and the anterior interposed nucleus (NIA). At this low magnification, the plexuses of nerve terminals in the nuclei appear as 'clouds' of fine dots representing individual varicosities. In the lateral cerebellar nucleus, the sector receiving labelled terminals is sharply demarcated from neighbouring tissue that does not receive labelled innervation, suggesting a detailed topographical organization of the PO projection to the lateral cerebellar nucleus. Bar = 250/tm. B. Photomontage of olivary axon in the border of the white matter and interposed nuclei of rat cerebellum which gives off a thin collateral toward an area of innervation. The branching point is indicated by an arrow. The thin collateral follows a tortous course (for a short distance it is obscured by an underlying thicker axon) and demonstrates several varicosities that may be the sites of synaptic interaction. Bar = 20/Lm. C. Darkfield micrograph of a sector of the lateral cerebellar nucleus in a sagittally sectioned specimen. Thicker olivocerebellar axons (arrow) run in the overlying white matter and some traverse the neuropil in the right part of the micrograph. These arriving olivary axons seem to give rise to the dense plexus of thin varicose terminal fibers, which extend over most of the illustrated neuropil. Bar = 20/tm. Van der Want et al. (1989a). (

such as the rostral parvocellular portion of the red nucleus and the ventrolateral nucleus of the thalamus, however, shows an increase rather than a decrease after lesions of the cerebellorubral and thalamic pathway. Several explanations were offered for this phenomenon, one of which involves the loss of a cholinergic, cerebellar efferent pathway that would facilitate glutamate release from corticothalamic or corticorubral terminals in these nuclei (Nieoullon and Dusticier, 1981; Nieoullon et al., 1984). The existence of cholinergic efferents among the projections of the cerebellar nuclei received support from observations by the same authors of a temporary decrease of ChAT activity in the red nucleus, especially in its rostral part, after lesions of the cerebellorubral fibers (Nieoullon and Dusticier, 1981). Moreover specific retrograde transport of [3H]choline in the cerebellorubral pathway to large perikarya in the anterior half of the lateral cerebellar nucleus of the rat was reported by Bernays et al. (1988) (Fig. 112) and to cerebellar nuclei in the cat by Stanton and Orr (1985). Evidence for the existence of cholinergic neurons in the cerebellar nuclei is controversial; ChAT was found to be absent in these cells (Kimura et al., 1981, cat; Armstrong et al., 1983, rat), but to be present in the large cells of Deiters' nucleus (Kimura et al., 1981). Ikeda et al. (1991) reported the presence of ChAT-immunoreactive neurons in the cerebellar nuclei of the cat; these cells give rise to both thalamic projections and nucleocortical collaterals. One group of small neurons in the rhesus monkey, which was described by Langer (1985) under the name of the basal interstitial nucleus (see Section 5.1 .), displays uniform and strong AChE activity. These small cells lie dispersed in the white matter of the flocculus and the nodulus and ventral to the dentate and the posterior interposed nucleus, in the roof of the fourth ventricle. The presence of strongly AChE-positive but ChAT-negative small cells in the white matter of the flocculus of the rat, that may correspond to Langer's interstitial nucleus, was noticed by Komei et al. (1983). The interstitial nucleus should be distinguished from the group y (Brodal and Pompeiano, 1957), which is a lateral extension of the superior vestibular nucleus, located ventral to the dentate nucleus. The cells of group y are slightly larger AChE-positive neurons. In accordance with other parts of the superior vestibular nucleus, group y contains a small number of GAD-positive cells, but most of its cells display aspartatelike immunoreactivity (Kumoi et al., 1987). 163

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Reports on the presence of other transmitters or transmitter-related substances have been published. Some neurons in the medial and posterior interposed nucleus of the opossum were immunoreactive with antibodies to CCK (King and Bishop, 1990, Fig. 116). Some somatostatin-like immunoreactive neurons were present in the medial part of the fastigial nucleus of the rat (Vincent et al., 1985). Occasional cells labelled with antibodies against conjugates of taurine were observed by Ottersen et al. (1988b) in rat cerebellum. Glycine-immunoreactivity was observed in large neurons of the ventral fastigial nucleus of the rat (Chen and Hillman, 1993b). PKC subtypes in neurons of the cerebellar nuclei have been specified by Kose et al. (1988), Shimohama et al. (1990), Huang et al. (1991), Merchenthaler et al. (1993), Garcia et al. (1993) and Chen and Hillman (1993a) (Table 1). 5.5. A F F E R E N T CONNECTIONS OF THE CEREBELLAR NUCLEI: P U R K I N J E CELL AXONS The Purkinje cell axons constitute the main afferent system of the cerebellar nuclei. Other afferents include collaterals of mossy and climbing fibers, a projection from the contralateral red nucleus (see Section 5.6. and 6.4.4.) and certain monoaminergic systems (see Section 3.8.). The terminals of the Purkinje cell axons in the cerebellar nuclei and the lateral vestibular nucleus contain pleomorphic synaptic vesicles, most of which are elliptical and are provided with Gray type II or an intermediate type of synapse. These terminals cover most of the soma and the large dendrites, extend on the spines and the small dendrites and are found as axo-axonal synapses on the initial segment. These terminals first were identified by antegrade axonal transport of [3H]-GABA in the rat (McGeer et al., 1975) and tritiated leucine in the cat (Walberg et al., 1976). Most of the boutons lining the surface of cells of the cerebellar nuclei contain GAD or GABA-like immunoreactivity (Saito et al., 1974; Wassef et al., 1986; Fonnum et al., 1970; Fonnum and Walberg, 1973; Houser et al., 1984; Roffler-Tarlov et al., 1979; Kumoi et al., 1987; Hawkes and Leclerc, 1986). Most if not all Purkinje cells are GABAergic (Chan-Palay, 1984; Mugnaini and Oertel, 1985), but with respect to other substances they constitute a heterogeneous population (see Sections 3.1.2. and 3.1.3.). The possibility of a differential distribution in the cerebellar nuclei of the terminals of chemically different populations of Purkinje cells has been studied for taurine and for the Purkinje cell specific antibody mabQ 113 (anti-Zebrin). For taurine the results were conflicting. The distribution of taurine in parasagittal bands of Purkinje cells in the cerebellar cortex was based upon the presence in these cells of the synthesizing enzyme of taurine CSADase (Chan-Palay et al., 1982a,b). No such a differential distribution was noticed, however, for taurine-like immunoreactivity in Purkinje cells (Madsen et al., 1985; Ottersen and Storm-Mathisen, 1987) or for taurine-containing terminals in the cerebellar nuclei (Ottersen et al., 1988b). Zebrin-immunoreactive and non-immunoreactive Purkinje cells are distributed in parallel longitudinal bands in the cortex of rat cerebellum (Hawkes et al., 1985) and distribute their axons to different parts of the cerebellar nuclei (Hawkes and Leclerc, 1986). Light microscopical observations showed that all Zebrin positive boutons on the soma and dendrites of large central nuclear cells contained GAD, and that most GADpositive boutons on individual cells either were Zebrin-positive or -negative. The two populations of Purkinje cells, therefore, terminate on different central nuclear cells. Zebrin-positive Purkinje cells of the vermis projected to the caudal part and a majority of Zebrin-negative Purkinje cells to the rostral part of the fastigial nucleus of the rat. 164

The cerebellum." chemoarchitecture and anatomy

Ch. I

Hawkes and Leclerc (1986) estimated that Zebrin-positive terminals made up the great majority of the GAD-positive boutons on the class of Zebrin receptive neurons. This suggests that intrinsic GABAergic terminals are few and that extracerebellar and intrinsic non-GABAergic connections account for at most 20% of the terminals on these cells. GABA receptors in the cerebellar nuclei were not systematically studied (see Section 3.7.). 5.6. EXTRACEREBELLAR AFFERENTS OF THE CEREBELLAR NUCLEI: COLLATERALS OF MOSSY AND CLIMBING FIBERS Extracerebellar afferents of the cerebellar nuclei consists of the collaterals of mossy and climbing fiber systems, that provide their excitatory drive (Eccles et al., 1967), the cholinergic (Section 3.10.), and the monoaminergic afferent systems. The existence of extracerebellar afferents to the central nuclei is known from antegrade axonal tracing experiments and was verified in ultrastructural studies. Injection of [3H]-leucine or other antegrade tracers in the inferior olive in the cat and the rat resulted in labelling over climbing fiber strips in the molecular layer and over parts of the cerebellar nuclei and the lateral vestibular nucleus (Courville, 1975; Groenewegen and Voogd, 1977; Groenewegen et al., 1979; Kawamura and Hashikawa, 1979; Balaban, 1984, 1988; Van der Want and Voogd, 1987; Van der Want et al., 1989a and b). The presence of projections of the inferior olive to the vestibular nuclei outside the lateral vestibular nucleus was denied by Groenewegen and Voogd (1977), but advocated by Balaban (1984, 1985, 1988) in experiments on rabbits. When the projection of the inferior olive to the fastigial nucleus was analysed with ultrastructural autoradiography of [3H]-leucine in the cat, the labelled boutons were predominantly found on small, distal dendrites, but never on somata. These terminals contain spherical vesicles and occasional dense core vesicles in an electron-lucent matrix and were provided with asymmetrical synapses (Van der Want and Voogd, 1987) (Fig. 115). The origin of these terminals as collaterals from olivocerebellar fibers terminating as climbing fibers in the cortex was first suggested by electrophysiological studies (Eccles et al., 1967) and by the observation of a strict topographical relation between the olivocerebellar climbing fiber zones and the termination of olivocerebellar fibers in the cerebellar nuclei (Groenewegen and Voogd, 1977). Direct proof of a collateral origin of olivonuclear fibers was provided by the retrograde transport to the central nuclei of [3H]-D-aspartate, injected in the cerebellar cortex of rats (Wiklund et al., 1984); the double-labelling studies in the cat with fluorogold implants in the cerebellar nuclei combined with injections of rhodamine-spheres in the cortex (Qvist, 1989b) and the observations of collaterals in antegrade tracing with the lectin from Phaseolus vulgaris in the rat (Van der Want et al., 1989a,b) (Fig. 113). Collaterals from olivocerebellar fibers generally terminate in the particular cerebellar nucleus that receives the axons of the Purkinje cells innervated by the same set of olivocerebellar fibers. It is not known whether all subdivisions of the inferior olive project to the cerebellar nuclei. The topographical distribution of the olivonuclear projections is reviewed in section 6.3.3. Some of the nuclei in the brainstem and the spinal cord, that give rise to mossy fibers terminating in the granular layer, also project to the cerebellar nuclei. Antegrade and retrograde axonal tracing experiments demonstrated projections in rat and cat from the spinal cord (Szentagothai in Eccles et al., 1967; Voogd, 1969; Matsushita and Ikeda, 1970; Robertson et al., 1983; Ikeda and Matsushita, 1973; Matsushita and Yaginuma, 1990, 1995), the lateral reticular nucleus and from the nucleus reticularis tegmenti pontis 165

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Fig. 114. EM autoradiogram of a spiny dendrite in the fastigial nucleus of the cat. A labelled mossy fiber terminal, originating from an injection of tritiated leucine into the nucleus reticularis tegmenti pontis, is densely filled with uniform spherical vesicles. Boutons with flattened vesicles form synaptic contacts on the same dendrite. Cat. Van der Want et al. (1987).

and adjacent regions of the pontine nuclei (Ktinzle, 1975; Russchen et al., 1976; ChanPalay et al., 1977; Matsushita and Ikeda, 1976; Eller and Chan-Palay, 1976; Martin et al., 1977; Ruggiero et al., 1977; Hoddevik, 1978; Dietrichs and Walberg, 1979a, 1987; Dietrichs, 1983b; Brodal et al., 1986; Gerrits and Voogd, 1987; Qvist, 1989a,b; Shinoda et al., 1992; Mihailoff, 1993). Other mossy fiber systems, such as the cuneocerebellar tract and the basal pontine nuclei provide only few or no collaterals to the cerebellar nuclei. Their distribution is reviewed in Section 6.4.4. Terminals in the fastigial nucleus of the cat originating from the reticular and the vestibular nuclei ranged widely in size and formed asymmetric synapses with small and large dendrites but not with somata. These boutons contained clear, spherical vesicles, and they occurred in two types, that differed in the aggregation of their vesicles (ChanPalay, 1977) which are present in equal numbers among the terminals from all sources (Van der Want et al., 1987) (Fig. 114). Morphologically these boutons are indistinguishable from the olivonuclear boutons that were described in Van der Want's companion study (Van der Want and Voogd, 1987) (Fig. 115). The differences in distribution between the climbing fiber and the different types of mossy fiber terminals in the rat and monkey lateral cerebellar nucleus reported by Chan-Palay (1973a, 1977), that were based on the similarity in morphology of these terminals in the cerebellar cortex and the nuclei, were not confirmed. According to the ultrastructural degeneration studies of Ikeda and Matsushita (1973) spinocerebellar fibers terminate both on dendrites and on somata. 166

The cerebellum." chemoarchitecture and anatomy

Ch. I

5.7. EXTRACEREBELLAR A F F E R E N T S OF THE CEREBELLAR NUCLEI: SEROTONINERGIC, N O R A D R E N E R G I C , D O P A M I N E R G I C AND PEPTIDERGIC PROJECTIONS Serotonin-like immunoreactivity resides in a fine network of varicose fibers in the neuropil of all cerebellar nuclei. This plexus is most dense in the hilar region of the lateral cerebellar nucleus of the rat (Takeuchi et al., 1982), among the small, intrinsic neurons of this region (Chan-Palay, 1977) and in the caudal and dorsal regions of the central nuclei of the opossum (Fig. 117) (Bishop et al., 1985). The noradrenergic innervation of the cerebellar nuclei was studied with histochemical fluorescence methods by H6kfelt and Fuxe (1969) in the rat and Landis et al. (1975) in mice and by Mugnaini and Dahl (1975) in the chicken and with selective uptake of [3H]noradrenalin (Chan-Palay, 1977) and dopamine-fl-hydroxylase immocytochemistry (Pasquier et al., 1980) in rats. The network of varicose noradrenergic fibers in the cerebellar nuclei is much less dense than the serotoninergic plexus and in the chicken a noradrenergic innervation of the central nuclei even is completely lacking (Mugnaini and Dahl, 1975). The possibility should be considered that this plexus mainly consists of passing fibers on their way to the cortex (Sachs et al., 1973). A heterogeneous population of terminals, containing large dense-core vesicles with a diameter of 900 A, were found to be labelled in the cerebellar nuclei after intraventricular infusions of 3H-serotonin in the rat. Few of these boutons show synaptic specializations (Chan-Palay, 1975, 1977). The ultrastructural morphology of nor-

Fig. 115. EM autoradiogram showing two climbing fiber boutons with spherical and pleomorphic vesicles, labelled from an injection with tritated leucine into the inferior olive make synaptic contact with a dendrite of a neuron of the fastigial nucleus of the cat. Van der Want and Voogd (1987).

167

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J. Voogd, D. Jaarsma and E. Marani

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adrenergic terminals in the cerebellar nuclei has not been studied. Ultrastructural studies of the central nuclei using antibodies against conjugates of serotonin or noradrenalin have not yet been reported. Neurons of the ventral tegmental area of the rat project to the cerebellar nuclei. These neurons do not contain tyrosine-hydroxylase and this projection, therefore, is nondopaminergic, contrary to the projection of the dopaminergic A10 group located in the same area, to the cerebellar cortex of the rat (Ikai et al., 1992). Peptides, such as enkephalin, corticotropin-releasing factor (CRF) and cholecystokinin (CCK) that occur in both mossy and climbing fibers(see Sections 6.3.2.2. and 6.4.3.) have been demonstrated in the cerebellar nuclei of several mammals. The distribution of these peptides in a plexus of beaded fibers in these nuclei is very similar the plexiform distribution of serotonin and noradrenalin, substances that do not occur in climbing or mossy fibers. Enkephalin-like activity was concentrated in fibers along the lateral and dorsal borders of the dentate nucleus and in the rostral part of the fastigial nucleus and in varicosities in all parts of the cerebellar nuclei of the opossum (King et al., 1987) (Fig. 188). Enkephalin-like activity in cat and rat was found in some mossy fiber rosettes, but not in climbing fibers (King et al., 1987); the cerebellar nuclei have not yet been studied in these species. CRF-like immunoreactivity was present in a diffuse plexus of beaded fibers in all cerebellar nuclei of the opossum (Cummings et al., 1989) and in a similar plexus with concentrations in the anterior interposed nucleus and the ventral dentate in the cat (Cummings, 1989). CCK-like immunoreactivity was present in a similar plexus 168

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J. Voogd, D. Jaarsma and E. Marani

of varicose fibers in the cerebellar nuclei of the opossum (King et al., 1986b; King and Bishop, 1990).

6. EFFERENT AND AFFERENT CONNECTIONS OF THE CEREBELLAR CORTEX: CORTICONUCLEAR, OLIVOCEREBELLAR AND M O S S Y FIBER C O N N E C T I O N S AND CYTOCHEMICAL MAPS

Morphological, embryological, physiological and cytochemical studies have revealed longitudinal, zonal patterns in the cerebellar cortex. The cytoarchitecture of the cerebellar cortex is uniform, without clear signs of a regional differentiation. Differences in the size of granule and Purkinje cells between vermis and hemisphere, with larger and less densely packed cells in the vermis, and smaller more densely packed cells in the hemisphere, were noticed by Lange (1982) and Drfige et al. (1986) for human and rat cerebellum. Local differences also exist for the Golgi and/or the unipolar brush cells, which are more closely packed in the flocculus and the vermis of the posterior lobe, than in the anterior vermis and the hemisphere. Maps showing local differences in density of the Purkinje cells have been published for the cerebellum of the turtle Pseudomys scripta elegans and the lizard Varanus exanthematicus (Gerrits and Voogd, 1973; Bangma et al., 1983). The cerebellum of these species is rather simple and consists of a single leaf. Three zones with a different density of the Purkinje cells, that also differ in their projection to the cerebellar and vestibular nuclei were distinguished. Similar density maps are not available for the mammalian cerebellum, probably because the curvature of the cortex makes this technically difficult. There are indications in the mammalian cerebellum for medio-lateral, zonally distributed differences in the size of the Purkinje cells (Chan-Palay et al. 1981; Voogd, 1989) and an even better case can be made for the existence of systematic differences in the size of the fibers taking their origin from different Purkinje cell zones (Fig. 118) (Voogd, 1964, 1967, 1969; Voogd and Bigar6, 1980). The mediolateral compartmentalization of the Purkinje cells and their axons is related to the zonal organization of the corticonuclear projection and was first recognized by Klimoff (1899). He also concluded from his Marchi experiments in the rabbit that the corticonuclear projections are uncrossed, each hemivermis projecting to the medial cerebellar nucleus and to the vestibular nuclei, and the hemisphere to the dentate/interposed complex. The differential projection of the left and right hemivermis to the medial nuclei of either side implies that a sudden transition in the corticonuclear projection, in this case across the midline, is perfectly compatible with the uniform structure of the adult cortex. During early, fetal stages of cerebellar development the cortex is not uniform and discontinuities in the anlage of the Purkinje cell layer are present that subdivide it into a number of bilateral symmetrical parasagittal zones or clusters (see Section 6.2.). The first to demonstrate the paired origin and the zonal pattern in the development of the cortex were Hayashi (1924) and Jakob (1928) who distinguished an intermediate zone (the 'pars intermedia', Zwischenstiick or B convolution) located between the anterior vermis and the hemisphere in the 3-4 month human fetus (Fig. 154). The anlage of the anterior vermis was paired and consisted of the two A convolutions. Its histogenesis was more advanced than in the pars intermedia. Development of the cortex in the hemisphere lagged far behind. Caudally the triangular intermediate zone extended into the posterior lobe in the region of the paramedian sulcus where it degenerated and disappeared. The dentate nucleus developed in the hemisphere, the emboliform nucleus 170

The cerebellum." chemoarchitecture and anatomy

Ch. I

Fig. 118. Myeloarchitectonic compartments of the ferret cerebellum. In A the thin fibers at the border of the A and B compartments (R) are shown. In B the small Purkinje cell fibers of the C2 compartment are flanked by larger Purkinje cell fibers of C, and C3. H~iggqvist stain. Bar = 100/lm. Voogd (1969). (anterior interposed nucleus, Hayashi 1924) and the globose nucleus (posterior interposed nucleus, Jakob, 1928) belonged to the pars intermedia and the medial nuclei occupied the white matter of the vermis. Jansen and Brodal (1940, 1942) studied the corticonuclear projection of the corpus cerebelli with the Marchi method in cat, rabbit and macaque. They confirmed the uncrossed projection of each hemivermis to the medial nucleus and the vestibular nuclei. In the hemisphere they distinguished an intermediate zone, that projects to Brunner's (1919) interpositus nucleus and a lateral zone, that is connected with the lateral nucleus. They noticed the correspondence between their three-zonal arrangement in the corti171

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J. Voogd, D. Jaarsma and E. Marani

conuclear projection of the anterior lobe and the corticogenetic zones of Hayashi and Jakob, but at the level of the cortex the exact borders of the three zones of Jansen and Brodal remained arbitrary. The paramedian sulcus, which is the border between vermis and pars intermedia, is often absent or indistinct and the border between the intermediate and lateral zones in Jansen and Brodal's scheme depends upon the arbitrary border between the interpositus and lateral nuclei. Moreover the longitudinal topology of their zones was defective. In the anterior lobe the orientation of the three zones was approximately perpendicular to the interlobular fissures, but when they were extrapolated to include the posterior lobe, the intermediate and lateral zones did not follow the curved axis of the folial chain of the hemisphere, and their common border cut through the centers of the folial loops of the ansiform lobule and the paraflocculus. Brodal's (1940; see also Jansen and Brodal, 1958) earlier studies of the olivocerebellar projection supported the distinction of the three zones in the anterior lobe. In the cat the dorsal accessory olive was found to project to the vermis and to the entire hemisphere of the anterior lobe. An additional projection of the rostral pole of the principal olive to the extreme lateral part of the anterior lobe could be observed in rabbit, monkey and man, but not in the cat. In the anterior lobe of rabbit and man, therefore, three zones can be distinguished on each side of the midline: the (hemi)vermis and the intermediate zone, that receive fibers from the dorsal accessory olive, and the lateral zone, that receives a projection of the principal olive and is absent in the cat. On the basis of these observations Jansen and Brodal (1940) called 'attention to the striking conformity in the arrangement of the corticonuclear and olivocerebellar projections, both systems apparently being arranged according to principles entirely different from those prevailing within the spinocerebellar and vestibulocerebellar projections'. The similarity in the organization of the olivocerebellar and corticonuclear projections remained one of the central concepts in the anatomy of the cerebellum that received ample support from later studies on longitudinal zonation. At the level of the cortex the borders between zones containing Purkinje cells that project to different target nuclei usually cannot be distinguished, but in the white matter the Purkinje axons from these zones collect in parasagittally oriented sheets that appear as white matter compartments in transverse sections. The sheets or compartments containing the Purkinje cell axons remain separated by narrow spaces. Voogd (1964, 1969) in cat and ferret, Marani (1982a, 1986) in the mouse and Feirabend and Voogd (1986) in the chicken traced these fiber compartments in myelin-stained (Hfiggqvist's method, 1936; Voogd and Feirabend, 1981) sections throughout the cerebellum. Large caliber fibers, that were identified as the axons of Purkinje cells, with an admixture of smaller fibers were present within the compartments and small myelinated fibers accumulate at their borders (Fig. 118). These borders usually are continuous with the borders between the subdivisions of the cerebellar nuclei and this configuration, therefore, made it possible to predict the longitudinal zonal organization in the corticonuclear projection (Figs 119 and 120). Corticonuclear projection zones are continuous from lobule to lobule and they are oriented perpendicular to the long axis of the folia: they follow the curved axis of the folial chains of vermis and hemisphere. The large caliber fibers within the compartments of the white matter could be identified as Purkinje cell axons with axonal tracing methods and immunohistochemistry with Purkinje cell-specific antibodies. They appear as discrete bundles, separated by narrow gaps with antibodies against cyclic GMP-dependent protein kinase (De Camilli et al., 1984) and calbindin-D28K (Paxinos, unpublished observations) in the rat and other species, with anti-Zebrin in cats and macaques (where Zebrin reacts with all Purkinje 172

The cerebellum." chemoarchitecture and anatomy

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of the zonal arrangement of the corticonuclear and olivocerebellar projection in the cat. The cerebellar nuclei are represented by circles. The diagram of the flattened inferior olive at the bottom of the figure is constructed according to Brodal (1940: Fig. 157). regions that are interconnected are indicated with the same symbols. The topography in the connections of the C] and C3 zones is indicated with open and filled diamonds. A - A zone; A N S I = ansiform lobule; B = cellgroup beta; B = B zone; C1-3 = C1-3 zones; D = D zones; D A O = dorsal accessory olive; dc = dorsal cap; dl = dorsal leaf of the PO; dmcc = dorsomedial cell column; F = fastigial nucleus; F L O C = flocculus; IA = anterior interposed nucleus; IP = posterior interposed nucleus; L = lateral cerebellar nucleus; LV - lateral vestibular nucleus; M A O = medial accessory olive; M E - medial extension of the ventral paraflocculus; P F L D - dorsal paraflocculus; PFLV = ventral paraflocculus; P M D = paramedian lobule; PO = principal nucleus of the inferior olive; SI - simple lobule; vest = vestibular nuclei; VI-X = lobules VI-X; vl - ventral leaf of the PO; vlo - ventrolateral outgrowth; X = X zone. Modified from Groenewegen et al. (1979).

173

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Fig. 120. Compartments in the white matter of the cerebellum of the cat. Drawings and reconstructions from H~iggqvist and AChE-stained sections. Compartments are indicated with different symbols. A-D. Graphical reconstructions of the rostral aspect of the anterior lobe (A) and the posterior lobe (B), the dorsal aspect (C) and the caudal aspect (D) of the cerebellum. Compare Fig. 98. E-G. Transverse sections. A = A compartment; ANS = ansiform lobule; A N T = anterior lobe; B = B compartment; C1-3 = C1-3 compartments; cr = restiform body; D(1,2) = D(1,2) compartments; F = fastigial lateral cerebellar nucleus; P F L = paraflocculus; P M D = paramedian lobule; SI = simple lobule; vest - vestibular nuclei; X - X compartment; III-IX = lobules III-IX. (.

cells) and with parvalbumin in the immature avian cerebellum (Braun et al., 1986) (Fig. 32). Monoamino oxidase in the white matter of the cerebellum of the chicken, is also restricted to the large caliber fiber areas (Feirabend, 1983; Marani, 1981). Systematic differences in the caliber of the Purkinje cell fibers were noticed in myeloarchitectonic studies in cat and ferret (Voogd, 1964, 1969; Voogd and Bigar6, 1980) and retrograde tracing of Purkinje cell axons (Voogd et al., 1991a). The small fibers located in the gaps between the bundles of Purkinje cell axons were called 'raphes' when they were positively stained with the H~iggqvist method (Fig.118; Voogd, 1964) and with AChE-histochemistry (Figs 122 and 127) (Hess and Voogd, 1986). Within the medullary core of the monkey cerebellum Hess and Voogd (1986) described fibers that stain densely for AChE and which are distributed in longitudinally oriented sheets, that appear as stripes in cross-sections (Fig. 127). These stripes in the white matter were aligned with AChE-rich bands, containing concentrations of reactive glomeruli in the granular layer, and narrow AChE-rich stripes in the molecular layer. A prominent midline band of AChE-rich axons flanked on each side by 5 or 6 ACHErich bands delineated a number of compartments in the cerebellar white matter. Within the white matter and the granular layer, cytochrome oxidase-rich axons and glomeruli were distributed in a longitudinally banded pattern, topographically identical to, but of lower contrast than, the banded distribution of ACHE. Both in the monkey and the cat the number and the disposition of the AChE-rich zones in the cerebellar white matter corresponded exactly with the raphe-like concentrations of small fibers at the borders of large fiber compartments in myelin-stained sections (Fig. 122). The highest amount of AChE was found in dense strips at the borders of the compartments; within the compartments the reactivity of the fibers for AChE differed. Compartments that contained large axons usually displayed very little AChE activity whereas the activity of AChE was high in compartments with smaller fibers. An explanation for the entire banded distribution of AChE in the molecular and granular layers and in the white matter as yet is not available, but it is clear that this distribution faithfully reflects the basic plan of the longitudinal zonation of the mammalian cerebellum. The possibility that zonally distributed differences in size and connections of the Purkinje cells are correlated with specific chemical properties of these cells was first raised by Marani (Marani and Voogd, 1977; Marani, 1981, 1982a; Marani, 1986) on the basis of the distribution of 5'-nucleotidase and acetylcholinesterase in the molecular layer and by Chan-Palay (1984) who reported a restricted distribution of certain peptides in subsets of Purkinje cells. More recently a complete pattern of alternating zones of immunoreactive and non-immunoreactive Purkinje cells was described by Hawkes and Leclerc (1986, 1987) with a Purkinje cell-specific antibody (anti Zebrin-I) in the rat and by Brochu et al. (1990) with anti-Zebrin II in the rat and other species (see Section 6.1.3.). The distribution of the Zebrin-positive Purkinje cells was very similar to the distribution of the enzyme 5'-nucleotidase in the molecular layer of certain rodents (Eisenman and Hawkes, 1989). 175

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The concept of the similarity in the organization of the corticonuclear and olivocerebellar projections, that dates from the work of Jansen and Brodal, received strong support from the observation that the distribution of the olivocerebellar fibers from certain subdivisions of the inferior olive was congruent with the parasagittal compartmentalization of the white matter (Voogd, 1969). The zonal termination of the climbing fibers could be visualized in autoradiograms of the olivocerebellar fibers using anterograde transport of [3H]leucine (Courville et al., 1974; Groenewegen and Voogd, 1977; Groenewegen et al., 1979). The olivocerebellar fibers located within a compartment were found to terminate both as climbing fibers on the Purkinje cells of the corresponding zone and on the cells of their cerebellar or vestibular target nucleus. The corticonuclear projection zones were experimentally verified in the antegrade degeneration and transport studies of Haines in primates and subprimates (see Haines et al., 1982 for a review) and Dietrichs in the cat (Dietrichs 1981a,b, 1983a; Dietrichs and Walberg, 1979a, 1980; Dietrichs et al., 1983b), and by retrograde labelling of the Purkinje cells and their axons from their target nuclei (Courville and Faraco-Cantin, 1976; Voogd and Bigar6, 1980; Balaban, 1984). A similar longitudinal pattern emerged from the electrophysiological studies of Oscarsson (1969, 1973, 1979, 1980; Oscarsson and Sj61und, 1974, 1977a,b,c; Andersson and Oscarsson, 1978a; Ekerot and Larson, 1979a,b; Ekerot et al., 1987, 1991a,b; Andersson and Eriksson 1981; Andersson and Nyqvist, 1983; Garwicz, 1992) and Armstrong et al. (1974) of the distribution of the climbing fibers in the cerebellar cortex. The electrophysiological identification of these climbing fiber zones rests on their laterality, their topography and on differences in latency between the different spino-olivocerebellar climbing fiber paths. These studies greatly extended our knowledge on the branching of the climbing fibers (Armstrong et al., 1973; Ekerot and Larson, 1982) and on the internal topography of the zones and lead to the distinction of the 'micro zone' as the smallest somatotopically defined unit engaged in the processing of information (Andersson and Oscarsson, 1978b; Garwicz, 1992). Direct proof of the congruence of the corticonuclear and olivocerebellar projections was obtained in the antegrade tracing studies of the Purkinje cells in electrophysiologically identified climbing fiber zones by Trott and Armstrong (1987a,b). The olivocerebellar and nucleo-olivary projections are reciprocally organized in the sense that a cerebellar target nucleus of one or more particular Purkinje cell zones is connected with the subdivision of the inferior olive that provides climbing fibers to the Purkinje cells of the same zone and a collateral projection to the target nucleus (see Ruigrok and Voogd, 1990, for a review of the literature). The zonal organization of the cerebellar cortex and the compartmental subdivision of the cerebellar white matter, therefore, are the manifestations of the modular organization of the output systems of the cerebellum. A module consists of a Purkinje cell zone, or a set of zones, its cerebellar or vestibular target nucleus with its GABAergic nucleo-olivary projection and its nonGABAergic output system and a sustaining olivocerebellar projection. Is the number of modules constant among the mammalian species and are they correlated with the different forms of chemoarchitectonic zonation in the cortex and the white matter? These questions only can be answered after a more thorough review of the corticonuclear projection (Sections 6.1.1., 6.1.2., 6.1.4. and 6.1.5.), the chemoarchitectonic evidence on zonation (Section 6.1.3.) and the olivocerebellar and nucleocerebellar projections (Section 6.3.) in different mammalian species. The existence of such a basic plan has been questioned (Boegman et al., 1988), not so much because the different maps for parasagittal zonation are mutually exclusive, but mainly because most 176

The cerebellum." chemoarchitecture and anatomy

Ch. I

of the studies on the topography of afferent and efferent connections were conducted in carnivores and primates, whereas the immunocytochemical studies, that revealed a finer grain in the parasagittal organization were mostly restricted to the rat. Evidence on the corticonuclear projection in cat and primates and its correlation with white matter compartments, mostly dating from the 1960s to the early '80s, will be discussed in Sections 6.1.1. and 6.1.2. The compartmentalization of the cerebellar cortex for Zebrin (Hawkes et al., 1985) and other markers, such as 5'-Nucleotidase (Scott, 1963), that was mostly studied in rodents, will be considered next (Section 6.1.3.). Correlations of the corticonuclear projection in the rat with these cytochemical maps are reviewed in Section 6.1.4. and efferent connections of the vestibulocerebellum in Section 6.1.5. Some regional differences in the development of the cerebellum, mainly concerning the transient, chemical heterogeneity of the Purkinje cells, are considered in Section 6.2. The inferior olive and the large body of anatomical and electrophysiological evidence on the olivocerebellar projection, that has contributed so much to the ideas on the zonal organization of the cerebellar cortex, are discussed in Section 6.3. 6.1. COMPARTMENTS AND CORTICONUCLEAR PROJECTION ZONES: CORRELATIONS WITH CYTOCHEMICAL MAPS 6.1.1. Corticonuclear projection zones in the cat: Correlation with white matter compartments and cytochemical zones

In the cerebellum of the cat at least 8 zones have been defined on the basis of the corticonuclear and olivocerebellar projections (Fig. 119). The corresponding white matter compartments have been delineated in myelin-stained or AChE-stained sections in ferrets (Voogd, 1967, 1969), cats (Voogd, 1964, 1989) and monkeys (Hess and Voogd, 1986; Voogd et al., 1987a,b; Voogd and Hess, 1989). AChE in the anterior vermis of the cerebellum of young cats is present in bands in the molecular layer (Marani and Voogd, 1977; Voogd and Bigar6, 1980) (Fig. 121). An AChE-positive band at the midline is separated by an AChE-negative area from a strongly reactive, paramedian band. At the lateral, sharp border of the paramedian band a narrow AChE-negative band is found, that merges into the hemisphere, which is uniformly AChE-positive. The bands diverge in the dorsal part of the anterior lobe and in the simple lobule. AChE in the molecular layer is present in parallel striations. The general direction of these striations corresponds to the orientation of the Purkinje cells and to the direction of the cortical zones. A similar, but less distinct pattern of AChE positive and negative zones is present in the lobules VIII-X of the caudal vermis. The AChE-stained material illustrated in this chapter differs from Marani and Voogd's (1977) original illustrations because it is derived from aldehyde-fixed tissue from the adult animal (Brown and Graybiel, 1983). Sections prepared in this way display the borders between the compartments in the white matter in addition to the banded pattern in the molecular layer. Three compartments can be distinguished on both sides of the midline in the region of the vermis of the anterior lobe: a medial A compartment, a lateral B compartment and a wedge-shaped X compartment in between A and B (Figs 120 and 122). Compartments A and B are present in all lobules of the anterior lobe, but an X compartment is only present in its dorsal part (lobules IV and V). Ventrally the fused AChE-positive borders of the X compartment continue in the lateral border of the fastigial nucleus. The fibers of the B compartment pass lateral to the fastigial nucleus and medial to the interposed nuclei to enter Deiters' lateral vestibular nucleus from dorsally. It is clear from a comparison of 177

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J. Voogd, D. Jaarsma and E. Marani

the AChE-positive borders between the compartments and the zonal distribution of AChE in the molecular layer, that the medial border of the B compartment is exactly in register with the lateral border of the first paramedian band of high AChE activity in the molecular layer. Consequently, the cortical X zone corresponds to the lateral part of this first paramedian band of AChE-activity, but its border towards the A zone cannot be recognized in the distribution of AChE in the molecular layer. The B zone corresponds to the second, AChE-negative strip of the molecular layer. Previously, the correspondence between the localization of AChE in the molecular layer and the corticonuclear projection has been studied by Brown and Graybiel (1983), but they failed to recognize the X zone. The molecular layer of the hemisphere of the anterior lobe is uniformly ACHEpositive. In the white matter of the hemisphere of the anterior lobe C1,C2,C3 and one or more D compartments can be delineated in AChE or HS.ggqvist-stained material (Figs 120 and 122). The C2 compartment is present in the dorsal part of the anterior lobe (lobules III, IV and V). More ventrally the C1 and C3 compartments are contiguous. The anterior interposed nucleus is located within the fused C1 and C3 compartments, the C2 compartment can be traced into the more caudally located posterior interposed nucleus. In H/iggqvist-stained sections of the cerebellum the fibers of the A,B,C1 and C3 compartments were larger than those of the X and C2 compartments. Staining for AChE generally is more intense in C2 and X than in the large fiber compartments. The AChE-banding pattern in the molecular layer of the anterior vermis and the white matter compartments of the anterior lobe continue, across the primary fissure, into the posterior lobe. A wide A compartment, flanked by diverging X and B compartments is present in the vermis of the simple lobule (Figs 119 and 120). The B compartment ends at the area without cortex in the center of the ansiform lobule, where it abuts on the pontocerebellar fibers of the cerebellar commissure that reach the surface at this point. It is not clear whether the X compartment continues from the simple lobule into lobule

Fig. 121. Zonal distribution of A C h E in the cerebellum of the cat. A. Whole mount preparation of the anterior lobe. B. Transverse section through the anterior lobe. H E M = hemisphere; m = midline AChE-positive band; L = parasagittal AChE-positive band; arrows = lateral border of L. Marani (1986).

178

The cerebellum." chemoarchitecture and anatomy

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Fig. 122. Photographs of white matter of the anterior lobe of the cerebellum of the cat. A. Borders of compartments and midline (m) are delineated by AChE-reactive raphes. The lateral border of the X compartment is in line with the lateral border of the parasagittal AChE-positive strip in the molecular layer; the molecular layer over the B compartment is AChE-negative. B. H~iggqvist-stain. Compartments A-D are delineated by dark stripes containing small calibre fibers. Same level as Fig. 120E.

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VII. Usually the entire white matter of lobule VII is considered to belong to the A compartment. 180

The cerebellum: chemoarchitecture and anatomy

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C1,C2,C3 and D compartments can be distinguished in the hemisphere of the simple lobule. C1 and C3 compartments are narrow or interrupted in the white matter of the ansiform lobule. A medial C2 and more laterally located D1 and D 2 compartments can be recognized in the white matter of this lobule. The complete set of compartments is present again in the white matter of the caudalmost folial rosette of the Crus II of the ansiform lobule (the ansula). From here these compartments pass into the paramedian lobule. C3 is present in the dorsal part of paramedian lobule only, and ends at the border with the pars copularis. C~ continues into the ventral most folia of the paramedian lobule as the most medial compartment of the hemisphere. C2 and the narrow, laterally located D1 and D2 compartments continue from the paramedian lobule into the dorsal paraflocculus (Figs 119 and 120). The dorsal and ventral paraflocculus contain C2, D1 and D 2 compartments. Concentrically arranged AChE-positive raphes, that stain strongly for ACHE, separate the compartments of the dorsal and ventral paraflocculus in the rostral part of the parafloccular loop (Figs 125 and 126). The rostral (dorsomedial) dentate nucleus is located within the D 2 compartment, the caudal (ventrolateral) dentate within the D~ compartment and the C2 compartment enters the lateral pole of the posterior interposed nucleus. Distinct compartments cannot be delineated in the white matter of the flocculus of the cat. The zonation in the corticonuclear projection that could be predicted from the disposition of the white matter compartments, was confirmed with retrograde labelling of Purkinje cells, after injections of tracers in the individual cerebellar nuclei of the cat (Courville and Faraco-Cantin, 1976; Voogd and Bigar6, 1980; Gibson et al., 1987). The Purkinje cells of the corticonuclear projection zones in the hemisphere follow the loops of the folial chain (Fig. 123, compare Fig. 98). Purkinje cells of the C1 and C3 zones can be retrogradely labelled from the anterior interposed nucleus. The C~ zone is broad in the ventral part of the anterior lobe and tapers more dorsally. The C3 zone is wide in the dorsal part of the anterior lobe, ventrally it is continuous with a narrow 'd2' zone 2 in the extreme lateral part of the anterior lobe. C1 and C3 are narrow in the ansiform lobule and reappear in the ansula and the paramedian lobule. They do not continue in the paraflocculus. The C2 zone was continuous from lobule III of the anterior lobe, through the entire hemisphere, into the flocculus. Its Purkinje cells were labelled from injections of the posterior interposed nucleus. Purkinje cells of the D zones occupied the extreme lateral part of the anterior lobe. A medial D1 and a lateral D 2 z o n e that projected to rostral and caudal parts of the lateral cerebellar nucleus, respectively, could be distinguished in the ansiform lobule, the paramedian lobule and the paraflocculus. The corticonuclear projection of the anterior lobe, the simple, ansiform and paramedian lobules was studied with anterograde transport of HRP by Dietrichs (198 l a) and Dietrichs and Walberg (1979a, 1980). Their observations on the projections of the A,B,C and D zones are essentially in accordance with the findings of Voogd and Bigar6 (1980). In their alternative nomenclature they shifted the zones one zone laterally, the presumed B zone projecting to the anterior interposed nucleus, and the C1 zone with the posterior interposed nucleus. This alternative nomenclature was discussed in the papers by Voogd and Bigar6 (1980), Haines et al. (1982) and Voogd et al. (1987a,b). The presence of C2, 2The term 'd 2' w a s applied to a narrow strip of climbing fiber-evoked potentials in the extreme lateral part of the anterior lobe by Ekerot and Larson (1979a, see Figs 171 and 175 ). The d 2 z o n e can be activated by the dorsal spino-olivo-cerebellar-climbing-fiber-pathand receives branches from climbing fibers which also innervate the lateral c3 zone (Ekerot and Larson, 1982).The use of the letter 'd' for this zone is misleading, because it neither projects to the lateral cerebellar nucleus or receives a projection from the principal olive. 181

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Fig. 124. Comparison of bands of AChE reaction product in the molecular layer of the anterior vermis of cat cerebellum and retrograde labelling of Purkinje cells in B and lateral A zones after an injection of HRP in the vestibular nuclei (A-C) and in the B and X zones after an injection in the lateral fastigial nucleus and the B compartment (D-F). Note different size of Purkinje cells in B and X zones. A = A zone; B = B zone; Deit = Deiters' nucleus; DV = descending vestibular nucleus; F = fastigial nucleus; IA = anterior interposed nucleus; MV = medial vestibular nucleus; X = X zone. I-V = lobules I-V. Voogd (1989). (

Dl and D2 zones in the paraflocculus of the cat is compatible with the anterograde tracing study of Dietrichs (1981 b). The corticonuclear projections of the electrophysiologically verified climbing fiber zones b, x, cl, c2, c3 and dl 3 w e r e determined for lobule V of the anterior lobe of the cat with anterograde tracing of [3H]leucine (Trott and Armstrong (1987a,b)). They were found to conform to the projections of the corresponding anatomical zones in the retrograde tracing studies of Voogd and Bigar6 (1980). The corticonuclear and -vestibular projection were correlated with the distribution of AChE for the anterior vermis of the cat by Brown and Graybiel (1983) and Voogd et al. (1991 a). Other studies of the projections of the anterior vermis to the fastigial nucleus and the vestibular nuclei, that have been known since Klimoff's (1899) study of the corticonuclear projection in the rabbit, did not distinguish between the zones in this region (Corvaja and Pompeiano, 1979; Dietrichs et al., 1983b; see Voogd 1964 and Brodal, 1974 for reviews of the older literature). Purkinje cells that project to the vestibular nuclei are located both in the A zone and in the B zone but not in the X zone (Fig. 124A-C). Purkinje cell axons of the A zone terminate in the fastigial nucleus, but some of these axons proceed to the vestibular nuclei, where they terminate at the border of Deiters' nucleus and the magnocellular medial vestibular nucleus (i.e. the ventral part of the lateral vestibular nucleus of Brodal and Pompeiano, 1957). The B zone projects to Deiters' nucleus (i.e. the dorsal part of the lateral vestibular nucleus). The projection of the A and B zones to different parts of the vestibular nuclei (Bigar6, 1980; Voogd and Bigar6, 1980; Voogd, 1989) was confirmed in the rabbit by Balaban (1984) and Epema (1990). These experiments substantiated the small projection of the A zone to Deiters' nucleus, as proposed by Andersson and Oscarsson (1978a). The wedge-shaped X zone in the dorsal part of the anterior lobe projects to the medial limb of the U-shaped nucleus at the junction of the fastigial and posterior interposed nuclei of the cat (Trott and Armstrong, 1987b). The Purkinje cells and their axons, when retrogradely labelled from this nucleus, are smaller than those of the A and B zones (Fig. 124D-F). The medial border of the retrogradely labelled B zone corresponds exactly with the lateral border of the parasagittal strip of high AChE-activity in the anterior vermis. The small Purkinje cells of the X-zone and the larger Purkinje cells with vestibular projections in the lateral A zone are located within the parasagittal AChE-positive strip. The B zone has a low content of AChE (Voogd, 1982). It was pointed out in a previous Section (5.3.) that the nucleocortical and corticonuclear projections in the cat are roughly reciprocal (Tolbert et al., 1978b; Gould, 1979; Dietrichs and Walberg, 1979a, 1980; Dietrichs and Walberg, 1985). Trott and Armstrong (1990) showed that nucleocortical projections to the electro-physiologically identified cl and c3 zones are scarce and that the major projection to the c2 zone of lobule V takes its origin from the posterior interposed nucleus.

30scarsson and co-workersadapted a modification of Voogd'snomenclature to indicate the electrophysiologically identified climbing fiber zones, using lower case letters instead of capitals (see Fig. 175). 183

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J. Voogd, D. Jaarsma and E. Marani

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PFL tI Fig. 125. White matter compartments C2, D1 and D2 in the paraflocculus of the cat in transverse, AChE-reacted sections. Note concentric arrangement of the compartments in the dorsal and the ventral paraflocculus. A caudalmost section; D = rostralmost section. ANS - ansiform lobule; brp = brachium pontis; C1-3 = C1-3 compartments; crest = restiform body; D - dentate nucleus; D (1,2)- D (1,2) compartments; F = fastigial nucleus; FLO = flocculus; IA - anterior interposed nucleus; IP = posterior interposed nucleus; PFLD - dorsal paraflocculus; PFLV- ventral paraflocculus; PMD = paramedian lobule. )

6.1.2. Compartments and corticonuclear projection zones in monkeys A l t h o u g h an analysis o f the c o m p a r t m e n t a l subdivision in the m o n k e y w o u l d be feasible f r o m serial HS, ggqvist-stained sections, the A C h E staining is m o r e distinct. A p r o m i n e n t c o n c e n t r a t i o n of A C h E - r i c h axons at the midline is flanked on each side of the anterior lobe by seven A C h E - r i c h strips, that delineate eight parasagittal c o m p a r t m e n t s in the cerebellar white m a t t e r (Fig. 127). Some of these c o m p a r t m e n t s are t o p o g r a p h i c a l l y related to certain cerebellar nuclei (Hess and Voogd, 1986; Voogd et al., 1987a,b). This is especially clear for two o f the c o m p a r t m e n t s , X and C2, b o t h of which have a high c o n t e n t of A C h E - p o s i t i v e fibres. The X c o m p a r t m e n t is n a r r o w a n d located in the white m a t t e r o f the vermis. It continues into the lateral, A C h E - r i c h b o r d e r z o n e o f the fastigial nucleus a n d the medial limb of the U - s h a p e d nucleus located between the caudal pole 184

The cerebellum." chemoarchitecture and anatomy

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25

of the fastigial and the posterior interposed nuclei (Fig. 105). In some sections the two AChE-rich strips that border the compartment, fuse with its content into a single, broad AChE-positive band. In the ventral part of the anterior lobe the X compartment is narrow or even absent. The X compartment extends caudally into the lobules VI and in the lateral parts of the lobules VII and VIII. The X compartment separates the A and B compartments of the anterior vermis. The A compartment includes the fastigial nucleus. In the posterior vermis it becomes wider and is subdivided into a medial A1 and a lateral A 2 compartment. The B compartment is present in the anterior lobe and the simple lobule. Ventrally it empties into the space between the fastigial and the anterior interposed nucleus to continue into the lateral vestibular nucleus. Caudally it diverges far laterally at the junction of the lobules VI and VII, where it ends at the area devoid of cortex in the centre of the ansiform lobule. The C2 compartment is clearly related to the posterior interposed nucleus. Rostrally it is located dorsal to the anterior interposed nucleus, in the intermediate part of the hemisphere. From the posterior interposed nucleus the C2 compartment extends dorsolaterally into the ansiform lobule, caudally into the paramedian lobule and ventrolaterally into the paraflocculus and the flocculus (Fig. 148). In the anterior lobe C2 is located between the C~ and C3 compartments. Ventrally Cl and C3 fuse and are related to the 185

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J. Voogd, D. Jaarsma and E. Marani

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Q Fig. 126. Zones and compartments in the paraflocculus of the ferret. C2, D~ and D2 zones indicated on surface of caudal (A) and rostral (C) aspect of the paraflocculus with different shadings. B and D. Reconstructions of the compartmental borders in the cerebellar white matter. ANS = ansiform lobule; FLO = flocculus; PFLD = dorsal paraflocculus; PFLV = ventral paraflocculus; PMD = paramedian lobule. Voogd (1969). Drawings by J. Tinkelenberg.

anterior interposed nucleus. Both C 1 and C 3 diverge laterally in the central part of the ansiform lobule and extend into the paramedian lobule. C1 ends in the last folium of the paramedian lobule, C3 stops in the dorsal part of this lobule. A major, lateral expanse of the white matter is related to the dentate nucleus. In the dorsal paraflocculus and the petrosal lobule this dentate-related white matter is subdivided into the D1 and D2 compartments (Fig. 148). D 1 is located next dorsal to the C2 compartment. D1 is related to the medial dentate nucleus. D2 occupies the dorsal part of the paraflocculus and is related to the lateral dentate nucleus. In the anterior lobe and the paramedian lobule 186

The cerebellum." chemoarchitecture and anatomy

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Fig. 12 7. Distribution of acetylcholinesterase at the borders of white matter compartments in the anterior lobe of Macaca fascicularis A-C3 = A-C3 zones; b c - brachium conjunctivum; b p - brachium pontis; F L O = flocculus; I A - anterior interposed nucleus; m - m i d l i n e ; P F L V - ventral paraflocculus.

a narrow compartment, lateral to C3, may represent D 1. The compartmental subdivision of the white matter of the ventral paraflocculus and the flocculus will be discussed in the Section on the vestibulo-cerebellum (6.1.5.). Our interpretation of this AChE pattern in the primate cerebellum generally is supported by data on the corticonuclear projection in primates and subprimates (see Haines et al., 1982 and Haines and Dietrichs, 1991 for reviews of the literature). Results from experiments on Galago, using silver impregnation of degenerated axons, showed that at least six zones corresponding to the A,B,CI__3 and D zones of carnivores, could be identified in the anterior lobe (Haines, 1977a; Haines and Rubertone, 1979) (Fig. 128). Lesions of the B zone in lobule V were associated with an additional projection to the 187

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J. Voogd, D. Jaarsma and E. Marani

most medial portions of the posterior interposed nucleus, suggesting the presence of an X zone in the vermis of this species (see also Haines and Dietrichs, 1991). Similar observations were made for the corticonuclear projection of the anterior lobe of Saimiri sciurus, where the X zone extended from lobule IV into VI. Purkinje cell axons of the X zone in the medial part of the posterior interposed nucleus do not overlap with the more centrally located terminations of the C2 zone in this nucleus (Haines et al., 1982; Haines and Dietrichs, 1991). Separate D1 and D 2 z o n e s with projections to rostrodorsal and centroventral regions of the lateral cerebellar nucleus, were distinguished in the anterior lobe in Saimiri (Haines et al., 1982). The cortico-vestibular projection of the anterior vermis was studied by Voogd et al. (1991 a) in Macaca fascicularis. The disposition of these Purkinje cells in the lateral A zone and the B zone of the anterior lobe and the simple lobule was similar to the cat. Retrogradely labelled Purkinje cell axons were located in the lateral A and the B compartments that could be delineated in adjacent AChE-stained sections. The X zone and compartment did not contain Purkinje cells with vestibular projections. Only few observations in primates are available on the corticonuclear projection of the posterior lobe. Haines and Whitworth (1978) and Haines and Patrick (1981) studied the projection of the paramedian lobule and the paraflocculus in the tree shrew (Tupaia glis). They concluded that Cl_3 and a D zone, with a similar topography and corticonuclear projection as in the cat, were present in the paramedian lobule of the tree shrew. The C2 and the D zone continued into the paraflocculus, where the D zone could be subdivided into D1 and D2 zones on the basis of its differential projection to the lateral cerebellar nucleus. The organization of the posterior vermis in primates (Haines, 1975a,b) will be dealt with in the Sections on the vestibular cerebellum (6.1.5.) and the olivocerebellar projection (6.3.3.3.).

VC

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Fig. 128. Diagrammatic representation of the corticonuclear projection of lobule V in Galago. There are at least six identifiable corticonuclear projection zones in the lobule V cortex. The vermis consists of zones A and B, the intermediate cortex of three zones C1 - C3 and the lateral cortex of a single D zone. f = flocculus; IC = intermediate cortex; LC = lateral cortex; lvn = lateral vestibular nucleus; 1-nia = lateral anterior interposed nucleus; m - nia = medial anterior interposed nucleus; m - nip = medial posterior interposed nucleus; nl = lateral cerebellar nucleus; nm = medial nucleus; vc - vermal cortex. Haines and R u b e r t o n e (1979)

188

The cerebellum." chemoarchitecture and anatomy

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The rough reciprocity of the corticonuclear and nucleocortical projections has been questioned in primates. In contrast to the observations in the cat Tolbert et al. (1978b), Tolbert and Bantli (1979) and Tolbert (1982) reported that all cerebellar nuclei projected to the vermal cortex in Macaca mulatta and that a major contingent came from the lateral cerebellar nucleus. Haines (1988) was unable to confirm these aberrant nucleocortical projections in Galago and suggested that some of Tolbert's injection sites in the lobules VI and VII of the caudal vermis may have extended in the neighbouring D zones. Haines (1989) found retrogradely labelled nucleocortical cells after injections of the C1, C3 and D zones to be relatively few and to be located in the corresponding anterior interposed and lateral cerebellar nuclei. The major nucleocortical projection in Saimiri takes its origin from the medial and posterior interposed nuclei, is bilateral and is directed at the A~ and C2 zones. This anisotropy in the nucleocortical projection in primates confirmed Trott and Armstrong's (1990) conclusions on the strong nucleocortical projection to the C2 zone in the cat. 6.1.3. Parasagittal zonation in the cerebellar cortex: Antigenic compartmentalization for Zebrin and other markers

The particulars of several Purkinje cell-specific markers that define parasagittal zones in adult rat cerebellum have been discussed in the first part of this chapter (see Section 3.1.2. for motilin, and the synthesizing enzyme of taurine (CSADase) and Section 3.1.8. for the zonal distribution of immunoreactive Purkinje cells in monkey and rat cerebellum, with the B.1 antibody of Ingram et al., 1985). CGRP, somatostatin and pseudocholinesterase (see Section 3.1.3.) are only present in zonally distributed bands of Purkinje cells during prenatal or early postnatal development. The complicated distribution of AChE was discussed in Sections 6.1.2. and 6.1.3. on the compartmental subdivision of the cerebellar white matter in cat and monkey. The distribution of AChE in the cerebellum of the rat is very similar (see Marani, 1986, Boegman et al., 1988; and Voogd, 1995 for reviews). Zebrin I and II (Section 3.1.8.) are the best-studied markers, that have been shown to be expressed by longitudinally organized subsets of Purkinje cells. Several other proteins and antigenic markers share the same or a very similar distribution as the Zebrins (the protein kinase C delta-isoform, Section 3.1.5.; the monoclonal antibody B30 of Stainier and Gilbert, 1989, Section 3.1.8.; the low affinity nerve growth factor receptor protein, Section 3.1.10.; and 5'-nucleotidase, Section 3.5.), whereas others only partially allign with the Zebrin pattern (HNK-1 antigen, Eisenman and Hawkes, 1993), or have a zonal distribution essentially complementary to the Zebrin pattern (cytochrome oxidase, Leclerc et al., 1990, P-path antigen, Leclerc et al., 1992, Section 3.1.8., and FAL in the Bergmann glia, Section 3.11.). Molecular markers of Purkinje cell heterogeneity include neurotransmitter receptors, such as the GABAB receptor (Section 3.7.2.), the muscarinic (m2) receptor (Section 3.10.2.), the dopamine D3 receptor (Section 3.8.) and the substance P receptor (Nakaya et al., 1994). It should be noted that the distribution of most markers, as identified in rodent species, may be different in other species or, as is the case for the muscarine m2 receptors, may be expressed in specific species only (Neustadt et al., 1988) (see Section 3.10.2.). An illustrative example of differential zonal distribution in closely related species was provided by Insel et al. (1994), who studied the distribution of vasopressin V~a receptors in the brain of different species of voles (microtine rodents). A complex distribution with alternating bands of high and low receptor density was observed in 189

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prairie voles. Instead, vasopressine Via receptors were confined to the nodulus in pine voles, and no receptors occurred in the cerebellar cortex of montane and meadow voles. The compartmentalization of motilin and taurine and the zonal patterns revealed by 5'-nucleotidase and Zebrin will be discussed in more detail. Chan-Palay (1984) described the distributions of Purkinje cells that react with antibodies against a conjugate of motilin (Chan-Palay et al., 1981; Nilaver et al., 1982) and against the synthesizing enzyme of taurine (L-cysteine sulfonic acid decarboxylase, CSADCase) (Chan-Palay et al., 1982a,b) (see also Magnusson et al., 1988, and Section 3.1.2.). Purkinje cells with motilin and/or GAD-like immunoreactivity together accounted for more than half of their total number. Purkinje cells that reacted with an antibody against motilin were most numerous in the flocculus and the paraflocculus. In the hemisphere they occurred in groups. They were fewer in the vermis, where they constituted a prominent midlineand two parasagittal bands in the lobules I-VI (Fig. 129). Motilin-immunoreactive Purkinje cells usually were larger than those containing only GAD. A majority of the Purkinje cells and many of the stellate and basket cells in the molecular layer reacted with the antibody against CSADC. These cells were distributed in a midline-band and 3 pairs of CSADC-positive and -negative bands on either side. The bands increased in width in the dorsal part of the anterior lobe (Fig. 130). In the vermis of the posterior lobe similar bands existed but were less distinct. Two CSADCpositive bands were present in the hemisphere. The flocculus and the paraflocculus contained the highest number of CSADC-positive Purkinje cells, but they were not zonally distributed. Chan-Palay's description of the zonal distribution of motilin and CSADC-immunoreactive Purkinje cells does not offer clues for a comparison with similar longitu-

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Fig. 129. Schematic drawing of the distribution ofmotilin-immunoreactive (M-i) Purkinje cells (open triangles) and glutamic acid decarboxylase-immunoreactive (GAD-i) Purkinje cells (filled circles) in a coronal section of rat cerebellum. M-i cells and GAD-i cells are both more concentrated in the flocculus and the paraflocculus than elsewhere. Both cell types occur in the vermis and participate in the formation of the sagittal microzones (arrows). M-i terminal axon projections in the deep cerebellar nuclei are heaviest in the dentate (D: left side) and GAD-i projections are heaviest in the lateral vestibular nucleus (LV: right side). ! = interposed nucleus; F = fastigial nucleus. Chan-Palay et al. (1981).

190

The cerebellum." chemoarchitecture and anatomy

Ch. I

III [anterior I. simple

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Fig. 130. Schematic summary of cysteine sulfinic acid decarboxylase (CSADCase)-positive sagittal microzones or bands in mouse cerebellum. The bands are clearest in the anterior lobe and the vermis, less sharply defined in the hemispheres (dense stipple), and most difficult to discern in the paraflocculus and flocculus (light stipple), because of intense CSADCase reactivity in most Purkinje cells. The dentate (D), interpositus (I), fastigial (F), and lateral vestibular nuclei (LVN) contain numerous CSADCase-positive cells. Chan-Palay et al. (1982b).

dinal patterns in the connections or in the distribution of other markers. The parasagittal zone of large, motilin-immunoreactive Purkinje cells may correspond to the B-zone, which, in the cat at least, contained larger Purkinje cells than the adjoining X zone. The majority of the terminals of the B-zone in Deiters' nucleus, however, contained GABA and did not react for motilin (Fig. 129). The distribution of 5'-nucleotidase (5"N) (Section 3.5.) in alternate longitudinal bands of high and low enzyme activity in the molecular layer of the cerebellar cortex of the mouse (Scott, 1963, 1964, 1965, 1967) was the first evidence for the biochemical compartmentalization of the cerebellar cortex. The pattern of 5'-N-positive and -negative zones is complete in the sense that it is present in all the lobules of vermis and hemisphere and unequivocal, because, in the mouse at least, the bands are clearly delineated (Marani, 1986). The 5'-N band pattern is very similar, if not identical, to the more recently described distribution of Purkinje cells in the rat, reacting with Purkinje cell-specific monoclonal antibodies to Zebrin-I (mabQ113) (Eisenman and Hawkes, 1989). The zonal distribution of 5'-N in the cerebellum of the mouse (see also Section 3.5.) was described in detail by Marani (1982a, 1986). A similar zonation of 5'-N was present in the cerebellum of the rat, the shrew (Marani, 1982a) and of Clethrionomys glarulus (Marani, 1982a). However, uniform and high levels of 5'-N without any indication of a longitudinal zonation characterize the molecular layer in some other rodents, the cat, some primates and in man (Scott, 1967; Marani, 1982a, 1986). The pattern in the rat is less distinct because a high background activity of 5'-N is present all over the molecular layer. This background activity disappears after long-standing lesions of the inferior olive (Marani, 1986). 191

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J. Voogd, D. Jaarsma and E. Marani

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Fig. 131. Reconstructions of the zonal distribution of 5'-nucleotidase (5'-N) in the molecular layer of the cerebellum of the mouse. Numbers without prefix indicate the nomenclature for the Y-N-positive bands of Marani (1982); P-numbers on the left side refer to the nomenclature for correspondingZebrin I-positivebands of Hawkes and Leclerc (1987). ANT = anterior lobe; FLO = flocculus; PFL = paraflocculus; II-X = lobules II-X. Marani (1982a, 1986). (

A midline band flanked by six, symmetrically disposed 5'-N-positive bands can be recognized in mice (Figs 58A, 131, 132). The bands were indicated by Marani (1982a) with the numbers 1--7 for the anterior lobe and the simple lobule and 11-17 for the posterior lobe. The bands in the anterior and posterior lobe are not necessarily continuous. Staining in the 5'-N-positive bands is rather uniform both in transverse and sagittal sections and cannot be assigned to specific structures in the molecular layer. There is an abrupt change in enzyme activity at the lateral border of the 5'-N-positive bands towards the next 5"N-negative zone; enzyme activity falls off more gradually at their medial borders. The 5'-N-positive bands share the distinctness of their lateral borders with the AChE-postive bands in the cerebellum of the cat and with the borders of the small fiber compartments (the 'raphes') towards the next lateral large fiber (Purkinje cell fiber) compartment in the cerebellar white matter. 5'-N-positive bands are narrow in the ventral part of the anterior lobe. They increase in width in the dorsal parts of the anterior lobe and the simple lobule and even more so in the rest of the posterior lobe, where the 5'-N-negative zones are reduced to narrow slits (Fig. 131). Staining in the medial bands 1-5 of the anterior lobe is heavier than in the more lateral bands 6 and 7 of the hemisphere of the anterior lobe. In the posterior lobe the intensity of the staining differs for the different bands. Heavy staining is found in the bands 11 and 13 and much less reaction product is present in the intermediate band 12 (Fig. 58A). The epitopes recognized by Hawkes' family of monoclonal antibodies known as the 'anti-Zebrins' are localized on Purkinje cells (see Section 3.1.8.). Zonal patterns that are identical or very similar to Zebrin I and II have been described for the distribution of 5'-nucleotidase (see above), the p75 low affinity nerve growth factor receptor protein in the rat (Section 3.1.10., Fig. 38), protein kinase C delta (Fig. 133) (see Section 3.1.5.) and the B30 antibody of Stainier and Gilbert (1989) (see Section 3.1.8.). Immunoreactivity in mouse Purkinje cells for an antibody against H N K is partially congruent with the Zebrin negative Purkinje cells, but Zebrin+/HNK+ Purkinje cells also exist (Hawkes, 1992; Eisenman and Hawkes, 1993). The similarity between the Zebrin pattern and the transient zonal patterns in the development of the Purkinje cell specific marker L7 is discussed in Section 6.2. Complementary staining patterns were described for cytochrome oxidase in Saimiri sciureus and rat by Leclerc et al. (1990), for the distribution of P-path-immunoreactive (Edwards et al., 1989, see below) Purkinje cells in the mouse by Hawkes (1992), Leclerc et al. (1992) and Edwards et al. (1994) and for 3-fucosyl-N-acetyl-lactosamine (CDts) in mouse Bergmann glia (Fig. 94) (Bartsch and Mai, 1991; Marani and Mai, 1992). According to Eisenman and Hawkes (1989) the 5'-N and Zebrin I zonal patterns in mouse cerebellum are congruent (Fig. 135). There are also similarities with respect to the mode and the time scale of their development. 5'-N in neonatal mice is distributed uniformly in the molecular layer and the first signs of a banded distribution cannot be discerned before postnatal day 14 (Hess and Hess, 1986). Zebrin I development in rats 193

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J. Voogd, D. Jaarsma and E. Marani

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Fig. 133. Distribution of protein kinase C delta-immunoreactive Purkinje cells in transverse section through the posterior lobe of rat cerebellum. This pattern is similar to the distribution of Zebrin. Bar - 0,5 mm. Chen and Hillman (1993a).

194

The cerebellum. chemoarchitecture and anatomy

Ch. I

also goes through a stage where all Purkinje cells express the epitope. The banded distribution appears relatively late, from postnatal day 15 onwards, when the immunoreactivity becomes suppressed in the future Zebrin I-negative zones (Leclerc et al., 1988). Differences between both patterns involve the distribution of the marker within the bands. Both for 5'-N and Zebrin I the intensity of the staining falls off in the more laterally located bands, but the sharp lateral borders and the differences in reactivity between the 5'-N-positive bands of the caudal vermis are not as clear with staining for Zebrin I. Moreover Zebrin I immunoreactivity extends into the Purkinje cell axons and compartments of Zebrin I-positive and -negative axons are present in the white matter that reflect the zonal distribution of the corresponding Purkinje cells. Zebrin I in rat cerebellum was compared to the distribution of AChE in the cerebellum of the rat (Boegman et al., 1988). These authors stressed the congruence of Zebrinpositive Purkinje cells with the accumulations of AChE in patches in the underlying granular layer. AChE in these patches is present in glomeruli, in certain Golgi cells and in other, unidentified components of the neuropil. Hawkes and Leclerc (1987) grouped the Zebrin I-positive Purkinje cells in a midline band (PI+) and seven symmetrically disposed parasagittal bands (P2+-PS+). Hawkes and Leclerc's (1987) numbering system for the Zebrin-positive and -negative bands in rat is indicated in Fig. 136. The P - bands of Zebrin-negative Purkinje cells bear the same number as the next medial P+ band. The pattern of Zebrin I-immunoreactive Purkinje cells is virtually identical in rat and mouse (Figs 139 and 140) (Eisenman and Hawkes, 1993). The numbering of 5'-nucleotidase-positive bands according to Marani (1986) and Hawkes and Leclerc's (1987) numbering system for the Zebrin-positive bands can be compared in Fig. 131 of the distribution of 5'-N in mouse cerebellum. The main features of the Zebrin pattern are the increase in width of the Zebrinpositive bands in the dorsal part of the anterior lobe, as compared to the ventral lobules I, II and III and the cortex in the bottom of the primary fissure (Figs. 139, 140, 143). The Zebrin-positive bands can be traced across the primary fissure in lobule VI that forms the rostral bank of the primary fissure. They also increase in width in dorsal lobule VI and fuse into an extensive, Zebrin-positive area that covers lobule VII and the adjoining ansiform lobule. Zebrin-negative bands reappear in caudal lobule VII and in the caudal folia of the Crus II and the paramedian lobule. The pattern is distinct in lobule VIII and the caudal copular portion of the paramedian lobule. Wide, Zebrinpositive separated by Zebrin-negative slits are present in lobule IX. Most Purkinje cells of lobule X are Zebrin-positive, although zonally distributed regions with higher and lower immunoreactivity can be recognized in the bottom of the postero-lateral fissure and in lobule X. The continuity of the Zebrin-positive zones is less clear than suggested by the published diagrams. Regions where the continuity of the zones cannot be assessed, include the lobule VII/ansiform lobule, where most Zebrin-positive Purkinje cells fuse into a single continuum, and the transitions between the lobules VII, VIII, IX and X in the bottom of the prepyramidal, secondary and posterolateral fissures. The P1 + band is narrow and consists of fused, bilateral portions. It is present in all lobules, fuses with the P2+ bands in the lobules VII and IX/X and is wider in lobule VIII. P2+ was identified in the anterior and posterior lobes, but its continuity cannot be established because it fuses with other P+ bands in lobule VII. P3+ is weakly immunoreactive in the anterior lobe, and looses its identity among the Zebrin-positive Purkinje cells of lobule VII. It reappears in caudal lobule VII and can be traced as a distinct band in lobule VIII. The apparent continuity of P3+ between the lobules VIII and IX may be false, a fusion of P3+ with P4+ into the P3+ band of lobule IX should 195

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Fig. 134. Diagrammatic reconstruction of the Zebrin II/P-'path staining pattern of the mouse cerebellar vermis. Areas that are Zebrin II positive are shaded: areas that are P-path positive are unshaded; regions that are double labelled are solid black. I-X, cerebellar lobules. Inset: double labelling for Zebrin II and P-path may be limited to the lateral half of the P3+ in lobule VIII. Redrawn from Leclerc et al. (1992).

be considered. P4+ is distinct in the anterior lobe and the simple lobule, fuses with the other P+ bands at the transition of lobule VII and the ansiform lobule, and reappears as a wide, Zebrin-positive strip at the border of vermis and hemisphere at the level of the Crus II and the dorsal paramedian lobule. P5+, P6+ and P7+ are present in the hemisphere of the anterior and posterior lobe, but cannot be traced as individual bands across the Zebrin-positive ansiform lobule. One or more of these P+ bands may continue in the Zebrin-positive, lateral pole of lobule IX. The relations of the P5+/P7+ bands with the uniformly Zebrin-positive Purkinje cells of the paraflocculus and the flocculus have not been established. 196

The cerebellum." chemoarchitecture and anatomy

A

Fig. 135. Photomicrographs of the ventral surface of the uvula of the cerebellum of the mouse in adjacent sections reacted with Zebrin I antibody (A) and for the presence of 5'-nucleotidase (B). Note the identical pattern of staining in both even though the borders of the 5'-nucleotidase staining (B) are less distinct. Q 113 = mabQ113, 5'N - 5'-nucleotidase. Eisenman and Hawkes (1989).

Short strips of Zebrin-positive Purkinje cells have been noticed between the P1 +, P2+ a n d P3+ b a n d s in the a n t e r i o r lobe a n d the simple lobule. Two n a r r o w strips of Zebrin-positive Purkinje cells were identified in the ansiform lobule, between P4+ and P5+. They were considered as bifurcations of these bands, that were indicated as P4+ a n d P 5 b + with the additional strips as P 4 b + a n d P5a+. The n a r r o w stretches of Zebrinpositive Purkinje cells in the dorsal vermis of the anterior lobe a n d the simple lobule, were considered as 'satellite bands'. They are i n c o n s t a n t a n d n o t necessarily bilaterally symmetrical. D o r 6 et al. (1990) identified alternating P+ a n d P - zones in the cerebellum of the grey

Fig. 136. The reconstruction of parasagittal bands of Zebrin I (mabQ113-immunoreactive) Purkinje cells in the adult rat cerebellar cortex as seen from the anterior (a) and posterior (b). The band pattern is based upon the serial reconstruction of nine complete and five partial cerebella from sections cut in the horizontal plane and four complete reconstructions from sections cut coronally. Bands P1 + through P7+ are labelled. Hawkes and Leclerc (1987). 197

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Fig. 137. Four frontal sections taken at different levels through the cerebellum of the adult Monodelphis immunoperoxidase stained by using antiZebrin II to reveal the compartmentalization of the vermis. The lobules are labelled with Roman numerals. The scale bar - 1,500/2m. Dot6 et al. (1990).

198

The cerebellum." chemoarchitecture and anatomy

Ch. I

opossum (Monodelphis domestica, Fig. 137). P l+ - P 3 + bands were identified in the anterior and posterior vermis. The precise identification of P2+ and P3+ was ambiguous in lobule V and different interpretations for the continuity of the reconstructed bands in the lobules VI-X were illustrated (Fig. 138). P4+ was located at the interface of vermis and hemisphere. It was located lateral to the area devoid of cortex at the border of lobule VII and the ansiform lobule, or was split in two Zebrin-positive bands, surrounding the cortexless area (compare Fig. 131 illustrating the 5'-N-positive band 15 in the cerebellum of the mouse) (Marani 1986). Zebrin I compartmentalization in Saimiri sciurus was studied by Leclerc et al. (1990). Both in the vermis and the hemispheres clusters of Zebrin I-immunoreactive Purkinje cells were separated by weakly stained Purkinje cell somata or unstained cells. Zebrinnegative bands, therefore, are less distinct than in rodents. P1 +, P2+ and P3+ bands are continuous from lobule to lobule and become narrower in the anterior lobe. P4+-P7+ bands were tentatively identified in the hemisphere, but not analysed in detail. A complementary histochemical zonation was detected for cytochrome oxidase, that was present in patches in the granular layer corresponding to the P - bands both in squirrel monkey and rat cerebellum. It is obvious from a comparison of the illustrations from the paper of Dor6 et al. (1990), showing the distribution of Zebrin I immunoreactivity in Purkinje cells and their axons and the zonation of AChE in monkey cerebellum, that the P2+ immunoreactivity in the anterior vermis corresponds to the X zone, and P 2 P2 P3

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Fig. 138. Three different interpretations of the adult Purkinje cell compartmentalization for Zebrin in the cerebellum of adult Monodelphis, illustrated in Fig. 136, are possible. A. The P2+ (dark grey) and P3 + (medium grey) bands are continuous and unbranched from lobule I to X. B. P2 + is continuous and unbranched, but a novel Zebrin II+ band is inserted between P1 and P2 in lobules VI to X (unshaded). C. P2+ bifurcates within lobule V to give two branches in lobules VI. Dor6 et al. (1990).

199

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Fig. 139. Drawings of three surface views of the mouse cerebellum, anterior, dorsal and posterior, showing the locations of the Zebrin+ bands of Purkinje cells. The Purkinje cell bands P1 +-P7 + are labelled in the dorsal view (for clarity, only the numerals have been used). Note that in the vermis of the posterior lobe the immunoreactive Purkinje cells form five to seven bands (posterior and dorsal views), whereas in lobules VII and VI all vermal Purkinje cells are immunoreactive (posterior and dorsal view). This pattern gradually changes in the anterior lobe to result in three to five very narrow immunoreactive bands (anterior view). In the hemispheres there are three major immunoreactive bands of Purkinje cells on either side (P5b +, P6 +, P7 +) plus two sub-bands in the paravermal area of the paramedian and ansiform lobules (P4b +, P5a +). Note too that the Purkinje cells are all Zebrin+ in the nodulus (lobule X, illustrated as indicated by arrows reflected out from the ventral surface of the cerebellum), the paraflocculus, and the flocculus. From Eisenman and Hawkes (1993).

200

The cerebellum." chemoarchitecture and anatomy

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to the B zone. Our preliminary observations on the distribution of Zebrin I-immunoreactivity in the cerebellum of macaques and cats suggested that all Purkinje cell somata in these species were Zebrin-positive and that compartmentalization was less distinct than in the reports on other mammalian species. Bands of P-path immunoreactive Purkinje cells alternate with zebrin II immunoreactive neurons in the cerebellum of the mouse (Leclerc et al., 1992) (Fig. 134). In the P3+ band in the anterior vermis, lobule VII, VIII and dorsal IX, the P4+ band in the lobules V and VIII and the P2+ band in dorsal lobule IX the two epitopes are colocalized. The B1 monoclonal antibody of Ingrain et al. (1985) also detects a subset of Purkinje cells in monkey cerebellum, but their distribution did not correspond to the distribution of Zebrin I in the squirrel monkey (Leclerc et al., 1990) or to AChE as reported by Hess and Voogd (1986). 6.1.4. The corticonuclear projection of the cerebellum of the rat: Correlations with zebrin-antigenic compartmentalization

The corticonuclear projection of the cerebellum of the rat recently was reviewed by Buisseret-Delmas and Angaut (1993) and Voogd (1995). It was studied with anterograde degeneration methods (Goodman et al., 1963; Haines and Koletar, 1979; Umetani et al., 1986) and anterograde tracing with [3H]leucine (Armstrong and Schild, 1978a,b). One interesting feature of these studies is that they document a projection from the hemisphere to the dorsolateral protuberance of the fastigial nucleus, that has never been observed in carnivores or primates. According to the experiments of Armstrong and Schild (1978b) this projection originates from the Crus II and the adjoining paramedian lobule, with smaller contributions of the Crus I and the copula pyramidis. Umetani et al. (1986) limited the projection to the dorsolateral protuberance to the cortex of the medial hemisphere of the lobules between the primary and prepyramidal fissures. The cortex of the copula pyramidis, caudal to the prepyramidal fissure was found to project to the medial part of the anterior interposed nucleus. The localization of Purkinje cells with circumscribed projections to single cerebellar nuclei was investigated with anterograde transport of WGA-HRP (Buisseret-Delmas, 1988a,b; Buisseret-Delmas and Angaut, 1993). Buisseret-Delmas (1988a,b; Fig. 141) distinguished A and B zones in the anterior vermis on the basis of their projection to the fastigial and the dorsal part of the lateral vestibular nucleus, and their afferent climbing fiber projections from the caudal medial and dorsal accessory olives. She distinguished the portion of the medial hemisphere between the primary and prepyramidal fissures, that projects to the dorsolateral protuberance, in the A zone as the lateral extension of the A zone (Fig. 142). The X zone was distinguished from the A zone by Buisseret-Delmas et al. (1993) and found to project to an area located at the junction of the fastigial and posterior interposed nuclei that they indicated as the 'interstitial cell groups'. C1, C2 and C3 zones projected to the interposed nucleus, and received their climbing fibers from the rostral half of the dorsal accessory olive (C1 and C3) and the rostral medial accessory olive (C2). C1 and C3 are interrupted in the Crus I and projected to medial and lateral portions of the interposed nucleus, the projection of C2 occupied its intermediate one third. Voogd's (1964, 1969) original definitions of the C zones, with C1 and C3 projecting to the anterior interposed nucleus, and C2 to the posterior interposed nucleus, therefore, were not retained in this study. Three D zones were identified by Buisseret-Delmas and Angaut (1989b) in the hemisphere of the cerebellum of the rat (Fig.141). The Do zone is unique for the rat, and 201

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Fig. 140. Computer drawing of the reconstruction of the Zebrin Purkinje cells bands in the unfolded adult C57/B6 mouse cerebellum. The drawing was from immunostained 40/~m thick coronal frozen sections. The continuity of the bands has been determined as best as possible. On the left and bottom are the scales in millimeters. The two axes have different magnifications. On the right are marked the approximate boundaries of the vermal lobules. The flocculus and paraflocculus are not illustrated. One place where the data are ambiguous is within lobule V-VI, where a large number of short bands more caudally are dramatically reduced to just three at the rostral limit. It is not clear whether the P2 + or P3 + bands extend through the anterior lobe vermis (see also Fig. 139). The reconstruction data from coronal sections were not suitable to resolve the issue, so the cerebellum has also been reconstructed from horizontal sections. The upper inset panel shows the data from such a reconstruction, equivalent to the region indicated by a rectangle on the main drawing (scale in millimeters). The preferred interpretation is that the P2 + compartment does not extend far into the anterior lobe vermis, and that the first lateral Zebrin+ band in lobules I-IV is continuous with P3 + (as indicated by continuous lines in the upper inset panel and as shown in the main drawing). The alternative hypothesis, that the first lateral Zebrin + band in lobules I-IV is continuous with P2 +, is shown schematically in the lower inset panel. Eisenman and Hawkes (1993). (

projects to the dorsolateral hump and receives a projection from the medial half of the ventral leaf of the principal olive. The Do zone is present in the anterior lobe and the lobulus simplex, Crus II and the paramedian lobule. It is interrupted at the level of the Crus I. The D1 and D 2 z o n e s occupy the lateral border of the hemisphere. D~ extends uninterruptedly from lobule III through the copula pyramidis. O 2 extends beyond the copula into the lateral parts of the vermal lobules IX and X and into the paraflocculus. The corticonuclear relations of the D1 and D 2 z o n e s of the rat are reversed with respect to their namesakes in the cat. D1 projects to the dorsal, magnocellular part of the lateral cerebellar nucleus; D2 to the ventral, parvicellular part of this nucleus. A detailed analysis is available for the projections of the pyramis and the copula pyramidis (Umetani and Tabuchi, 1988) (Fig. 153) and lobule VIIb with the rostral paramedian lobule (Umetani, 1989) (Fig. 152). The C1 zone, with a projection to the anterior interposed nucleus, was lacking in the paramedian lobule and wide in the medial cortex of the copula. C2 was present in both lobules and C3 was limited to the rostral half of the paramedian lobule. D1 (corresponding to Do of Buisseret-Delmas and Angaut, 1989b) projected to the dorsolateral hump and D 2 (corresponding to both their D1 and D 2 zones) to the lateral cerebellar nucleus. The topography of the C zones in the rat, therefore, corresponds to the situation in cat and monkey, with the exception of the projection of a zone in the medial hemisphere to the dorsolateral protuberance of the fastigial nucleus, the presence of an additional Do zone and the reversal in the corticonuclear projection of D1 and D 2. This correspondence, however, does not explain the pattern of Zebrin-positive and -negative zones in the same region. The projections of the cortex to the fastigial and vestibular nuclei were correlated with the Zebrin immunoreactive Purkinje cell zones, using double staining with an anti-zebrin antibody and cobalt-stabilization of the retrograde labelled Purkinje cells on the same section. Purkinje cells in the A and B zones were retrogradely labelled from injections of WGA-HRP in Deiters' nucleus (Voogd et al., 1991b). In the anterior lobe and the simple lobule they were located in the zebrin-negative P1- and the lateral P2- zones (Fig. 143, left). The labelled Purkinje cells in P2- (i.e. in the B zone) were bordered on their medial side by P2+. Zebrin-positive 'satellite' bands bordered the labelled Purkinje cells in the lateral P1- zone on their medial side. WGA-HRP injections at the junction of the fastigial and posterior interposed nucleus labelled Purkinje cells of P2+ and the satellite bands between PI+ and P2+ (Fig. 143, right). Injections of the dorsolateral 203

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protuberance of the fastigial nucleus labelled Purkinje cells located between the primary and prepyramidal fissures overlapping with and located in between the P4b+ and P5a+ zones (Fig. 144). Zebrin-positive and -negative zones in the anterior vermis, therefore, approximately correspond with the corticonuclear projection zones A, X and B that were known from studies in other species than the rat. The projection of the Zebrinpositive Purkinje cells of the X (P2+) zone to the junction of the fastigial and posterior interposed nucleus is in accordance with the tendency of Zebrin-positive Purkinje cells to project to the caudal pole of the fastigial nucleus (Hawkes and Leclerc, 1986). The P4b+, P4b- and P5a+ zones corresponded with the zone projecting to the dorsolateral protuberance. In the simple lobule this zone was located immediately lateral to the B zone. As a consequence the fused P4b+ and P5a+ zones continue, rostral to the primary fissure, as the P3+ zone of the anterior lobe. The topography of the C and D zones in the rat has not yet been correlated with the Zebrin pattern, but it seems likely that C1, C3 and D (Do and D2 zones of Buisseret-Delmas and Angaut, 1989b), projecting to the anterior interposed and the rostral dentate nucleus, will prove to correspond to Zebrinnegative zones and that the C2 and D1 zones, that project to the posterior interposed and the caudal dentate, will be Zebrin-positive. Figure 145 depicts a diagram illustrating this hypothesis.

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Fig. 143. Comparison between Purkinje cells of the A and B zone (open circles) retrogradely labelled from the vestibular nuclei (left side) and retrograde labelling of Purkinje cells of the X zone after an injection of WGA-HRP in the transitional region of the fastigial and posterior interposed nucleus the rat (right side). Purkinje cells of the X zone occupy the Zebrin-positive P2+ zone; labelled Purkinje cells of the lateral A zone and the B zone are located in the Zebrin-negative P1- and P2- zones. Graphical reconstructions of transverse sections double labelled for HRP reaction product and Zebrin I immunocytochemistry.COP = copula pyramidis; CrI and II = crus I and II of the paramedian lobule; PMD = paramedian lobule; S1 = simple lobule; I-X = lobules I-X. Voogd et al. (1991b). (,

6.1.5. The corticovestibular and corticonuclear projections of the flocculus and the caudal vermis. Correlations with cytochemical zones and compartments Larsell (1934) subdivided the cerebellum in a somesthetic corpus cerebelli and the vestibulocerebellum. The compartmentalization of the corpus cerebelli was considered in the previous sections. The vestibulocerebellum of Larsell consists of the flocculus and the nodulus, the caudalmost lobules of the folial chains of vermis and hemisphere. They are separated from the corpus cerebelli by the posterolateral fissure, one of the earliest fissures to appear during ontogeny. In primitive mammals a narrow band of cortex along the attachment of the roof plate of the fourth ventricle interconnect the nodule and the flocculus. The modular organization and the characteristic afferent and efferent connections of the vestibulocerebellum extend, beyond the posterolateral fissure into the ventral part of lobule IX and in the adjacent cortex of the ventral paraflocculus which, therefore, should be included in it. The distinction between the corpus cerebelli and the vestibulo-cerebellum is clearly revealed by calretinin. The rather uniform staining of granule cells and parallel fibers in the corpus cerebelli suddenly stops at the borders of dorsal and ventral lobule IX and of the paraflocculus with the flocculus. In the vestibulocerebellum the unipolar brush cells are heavily stained on a lightly stained background (Floris et al., 1994) (Fig. 146). Different zonally distributed Purkinje cell markers, such as Zebrin I (Hawkes and Leclerc, 1987), motilin, taurine (Chan-Palay, 1984) and somatostatin (Villar et al., 1989) occur more uniformly in the flocculus, and the paraflocculus. Most Purkinje cells of the flocculus and the paraflocculus are Zebrin-positive and no banding has been observed with immunocytochemical methods. Compartmentally distributed differences in fiber size were never observed in the flocculus of carnivores, but compartments can be delineated in the white matter of the flocculus in rabbits and in old and new world monkeys, both with a myelin (the HS.ggqvist) stain and AChE histochemistry. In the rabbit (Van der Steen et al., 1991, 1994; Tan et al. 1992, 1995a,b,c) and the monkey (Macaca fascicularis, Hess and Voogd, 1986; Voogd et al. 1987a,b) 5, respectively 4 compartments can be recognized in the flocculus (Figs 147, 148 and 149). The medialmost compartment 4 of the rabbit flocculus is narrow and located at the border of the middle cerebellar peduncle (Fig. 147a,d). Compartments 1 and 3 are relatively rich in AChE and fuse in the dorsal and caudal white matter of the flocculus. The compartment 2 is poor in AChE and contains large Purkinje cell fibers in the myelin-stained sections. The fifth, most lateral compartment is the caudal extension of the C2 compartment of the paraflocculus. The compartments 2-5 continue, across the posterolateral fissure into the paraflocculus. In the first folium of the ventral paraflocculus of the rabbit (folium p of Yamamoto and Shimoyama, 1977), the compartments 1 and 3 enclose the dorsal tip of compartment 2 (Fig. 147b,c). Four compartments 207

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Fig. 144. Localization of retrogradely labelled Purkinje cells (open circles) from an injection of WGA-HRP in the dorsolateral protuberance of the medial cerebellar nucleus (left) and of anterograde transport of Phaseolus vulgaris leucaglutinin (PhaL) in climbing fibers (stripes) from the medial portion of the MAO (tectorecipient area), with respect to bands of Zebrin-I labelled Purkinje cells in the rat. Reconstructions from sections double-stained for Zebrin and WGA-HRP or PhaL. Numbering of Zebrin-immuno-reactive Purkinje cell zones according to Hawkes and Leclerc (1987). COP = copula pyramidis; CrI(II)= crus I(II) of the ansiform lobule; PMD =paramedian lobule; SI = simple lobule; I-X = lobules I-X. Voogd et al. (199 lb). (

extend over the entire ventral paraflocculus of Larsell (1970) in old and new world monkeys. The most lateral compartment continues as the C2 compartment in the petrosal lobule (Fig. 148b,c), the medial three compartments do not extend beyond the narrow junction of the cortex of the ventral paraflocculus with the petrosal cortex. The narrow, medialmost compartment 4 of the rabbit flocculus was not recognized in the primate cerebellum. The compartments 1, 2 and 3 of the monkey flocculus, correspond to their namesakes in the rabbit. Fibers from the white matter of the flocculus and the adjoining folia of the ventral paraflocculus collect in a compact bundle: the floccular peduncle. The peduncle is applied to the ventral surface of the lateral parvocellular cerebellar nucleus and arches over the restiform body (Fig. 147c,f). Its fibers surround the ventral limb of the lateral cerebellar nucleus as a capsule in rabbit and cat. Medium sized cells of the dorsal group y are located between the fibers of the peduncle in these species. Smaller cells in a dense AChE-positive neuropil accumulate, as the ventral group y, between the peduncle and the cochlear nuclei with the restiform body and, more medially, at the entrance of the peduncle in the vestibular nuclei. In monkeys the group y is compact and situated as an enlongated mass in the floccular peduncle. More caudally, small AChE-positive cells of Langer's (1985) basal interstitial nucleus extend into the lateral compartments of the flocculus and in the AChE-positive raphes between them. Some fibers of the compartments 1 and 2 of the rabbit flocculus arch through the dentate nucleus, to join the floccular peduncle at the lateral border of the vestibular nuclei. These 'arciform' fibers have been found in cat, rat, rabbit and Galago but seem to be absent in primates (Langer et al., 1985b). The white matter compartments of the flocculus do not simply continue as components of the floccular peduncle, but a reorganization takes place, that directs Purkinje cell axons from the compartments 2 and 4 to the medial vestibular nucleus and of the compartments 1 and 3 to the superior vestibular nucleus. The C2 compartment does not contribute to the floccular peduncle, but leads its fibers towards the posterior interposed nucleus. In monkeys the equivalent of compartment 1 of the rabbit extends in the roof of the fourth ventricle, along the basal interstitial nucleus of the cerebellum (Fig. 148). The corticonuclear and corticovestibular projections of the flocculus were studied by Dow (1936), Bernard (1987) in rat, Voogd (1964), Angaut and Brodal (1967), Sato et al., 1982a and b) in cat, Dow (1938), Langer et al. (1985b) in primates and Yamamoto and Shimoyama (1977), Yamamoto (1978), De Zeeuw et al. (1994a) and Tan et al. (1995c) in rabbit. The differential projection of the Purkinje cell zones of the flocculus was discovered by Yamamoto and Shimoyama (1977). The precise correspondence of the corticovestibular and group y projections of the rabbit flocculus with the ACHEcompartmentalization of its white matter and its functional correlates were discussed by Van der Steen et al. (1994), De Zeeuw et al. (1994a) and Tan et al. (1995a,b) (Fig. 188). A projection of the flocculus to the nucleus prepositus hypoglossi has been dis209

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The cerebellum." chemoarchitecture and anatomy

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Fig. 145. Diagram of the Zebrin-antigenic zones (left) and the corticonuclear projection (right) in the rat. The diagram of the corticonuclear projection is based upon data from Umetani et al. (1986); Umetani (1989), Bernard (1987); Buisseret-Delmas and Angaut (1989b, 1993) and Voogd et al. (1991b). There is a good correspondance between the corticonuclear projection zones A, X and B of the anterior vermis, and for the projection of the lateral extension of the A zone of Buisseret-Delmas (1988a) and the Zebrin pattern. For other zones this correspondance is conjectural or absent. The corticonuclear projection of the region between vermal lobule VIc and VII and the lateral A zone is not known. The corticonuclear projection zones are indicated with the same symbols as their target nuclei in the bottom diagram of the cerebellar nuclei. Regions projecting to the vestibular nuclei (a lateral strip of the A zone, the B zone and the caudal vermis) are indicated with open circles. 1-7 - Zebrin antigenic zones P1-P7" A - zone A; A~ - lateral extension of the zone of Buisseret-Delmas (1988a); B - zone B; C1-3 - zones C1-C3; CrI - crus I of the ansiform lobule; CrII - crus I! of the ansiform lobule; D - zone D; Do =one Do of Buisseret-Delmas and Angaut (1989); DLH - dorsolateral hump; DLP = dorsolateral protuberance of the medial nucleus; DMC - dorsomedial crest; FLO - flocculus; IA - anterior interposed nucleus; I P - posterior interposed nucleus; L - lateral cerebellar nucleus; MM = medial part of medial nucleus; N C - caudomedial part of the medial nucleus; P F L - paraflocculus; PMD = paramedian lobule; SI - simple lopule; tM/IP - transitional region of the medial nucleus and posterior interposed nucleus (interstitial cell groups of Buisseret-Delmas et al. 1993); X - zone X; I-X- lobules I-X. (

chemistry have been p r o p o s e d for the caudal vermis of the cat (Voogd, 1964, 1969) and the m o n k e y (Voogd et al., 1987a,b, Tan et al., 1995b). The caudal vermis projects to the fastigial nucleus, but also to the (posterior) interposed nucleus (Bigar6, 1980, cat; A r m s t r o n g and Schild, 1978a; Bernard, 1987; U m e t a n i and Tabuchi, 1988; Tabuchi et al., 1989, rat) a n d the ventral, parvocellular p o r t i o n o f the lateral cerebellar nucleus (Van R o s s u m , 1969; Wylie et al., 1994, rabbit; Haines, 1977a Galago; Bernard, 1987; Tabuchi et al., 1989, rat) a n d the vestibular nuclei including the g r o u p y. L o b u l e VIII of the caudal vermis gives rise to a small projection to the dorsal part of the lateral vestibular nucleus (i.e. Deiters' nucleus), but other corticovestibular connections o f the caudal vermis and the flocculus avoid Deiters' nucleus and terminate in other subdivisions of the vestibular nuclei. Corticovestibular fibers from the lobules X and IX of the caudal vermis were studied by D o w (1936), B e r n a r d (1987), Tabuchi et al. (1989) in rat, Voogd (1964), A n g a u t and Brodal (1967) in cat and D o w (1938), Haines (1977a) in primates. A detailed study of the corticonuclear and corticovestibular projections was m a d e of the ventral face of lobule X of the rabbit cerebellum by Wylie et al. (1994). Their findings are s u m m a r i z e d in Fig. 150 and can be correlated with the c o m p a r t m e n t a l A C h E - s u b d i v i s i o n of the white m a t t e r of lobule X by Tan et al. (1995b) and the a n a t o m i c a l and electrophysiological analysis of the olivocerebellar projection to this lobule by K a t a y a m a and N i s i m a r u (1988), B a l a b a n and H e n r y (1988) and K a n o et al. (1990, 1991). Corticovestibular fibers f r o m the flocculus and the lobules X and IX of the caudal vermis terminate in roughly c o m p l e m e n t a r y areas of the vestibular nuclei (Fig. 151) (Dow, 1936, 1938; Voogd, 1964; A n g a u t a n d Brodal, 1967; Haines, 1977a; L a n g e r et al. 1985b; Bernard, 1987). The caudal vermis o f the cat usually was considered to belong to the A zone, that projects to the fastigial nucleus. A c c o r d i n g to Bigar6 (1980) (Fig. 123) and Voogd and Bigar6 (1980) the entire lobule VII and the medial two-thirds of the lobules VIII, IX and X are connected with this nucleus. Courville and D i a k e w (1976) d e m o n s t r a t e d the sequential representation of the lobules VI-VIII in the fastigial nucleus of the cat with lobule VII projecting to the tail of the nucleus and lobules VI and VIII to m o r e r o s t r o d o r s a l and ventral parts respectively. A similar connection o f lobule VII to the

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tail of the fastigial nucleus was demonstrated by Yamada and Noda (1987) for the macaque monkey. Purkinje cells in the medial two-thirds of lobule IX of the cat also project to the medial and descending vestibular nuclei; those destined for the descending vestibular nucleus are concentrated near the midline and at the lateral border of the A zone (Matsushita and Wang, 1986). The lateral one-third of lobule IX is connected with the posterior interposed nucleus (Bigar6, 1980). A complicated pattern of Purkinje cell zones and patches was reported for the organization of the projection of lobule X to the superior, medial and descending vestibular nuclei in the cat (Shojaku et al., 1987). The zonal organization of the efferent connections of the caudal vermis in the rabbit is quite complex, with discrete zones in the lobules IX and X projecting to the fastigial, descending, superior and medial vestibular nuclei, and lateral zones connected to the interposed and different subdivisions of the lateral cerebellar nucleus (van Rossum, 212

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$ Fig. 148. Compartmentation in AChE-stained transverse sections of the flocculus, the ventral paraflocculus and the petrosal lobule and the caudal hemisphere of Saimiri sciureus. Note AChE stained borders of the compartments and the differential staining in the molecular layer; AChE-positive cells of the basal interstitial nucleus of Langer are located along the borders of the compartments C2, 1 and 2 in (D) and (E). Group y is located within the floccular peduncle in (E). ANS = ansiform lobule; br.p = brachium pontis; c.rest = restiform body; C1-3 = C21-3 compartments; D = dentate nucleus; D1,2 = D1,2 compartments; F = fastgial nucleus; fis.post.lat = posterolateral fissure; FLOC = flocculus; IA = anterior interposed nucleus; IP = posterior interposed nucleus; lb.petr = petrosal lobule; PFLD = dorsal paraflocculus; PFLV = ventral paraflocculus. Courtesy of Dr. D.T. Hess.

1969). Some sort of zonal organization in the corticovestibular projection of these lobules in the rabbit also was reported by Balaban (1984), Epema et al. (1985) and Epema (1990). 214

The cerebellum." chemoarchitecture and anatomy

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More detailed, but still incomplete accounts of the corticonuclear projection of the caudal vermis are available for the rat. The terminal fields of the different lobules in the fastigial nucleus of the rat are organized in a circumferential manner (Armstrong et al., 1978a; Haines and Koletar, 1979) with more overlap between the lobules than has been reported for the cat (Courville and Diakew, 1976). PS.fillysaho et al. (1990) analysed the projections of the lobules VI-VIII to the fastigial nucleus in more detail in the rat. The terminal field of lobule Via is located in the middle subdivision of the fastigial nucleus, lobules VIb,c, VII and VII project both to the caudomedial and middle subdivisions, with the terminal field of lobule VIII reaching most rostrally. Terminations of these lobules in the dorsolateral protuberance are scarce or absent. A subdivision of lobule VII into a medial region, projecting to the caudomedial subdivision of the fastigial nucleus and of a lateral region projecting to the middle subdivision also was proposed by Umetani (1989) (Fig. 152). This medial region may correspond to the tecto-olivorecipient zone of Akaike (1986b, 1992) (Fig. 184). The lateral zone of lobule VII, that separates the tecto-olivo-recipient zone from the lateral extension of the A zone, with its projection to the dorsolateral protuberance of the fastigial nucleus may be identical to the X zone. 215

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Fig. 149. Diagrams of cerebellar cortex, showing the zonal configuration of the paraflocculus and the flocculus in rabbit, monkey and cat, based upon the compartmental subdivision of the paraflocculus and the flocculus in monkey and rabbit (Figs 147 and 148) and the olivocerebellar projection to the flocculus in cat (Voogd, 1964; Gerrits and Voogd, 1982) and rabbit (Tan et al., 1995a,b). Ca, D1 and D2 zones can be distinguished in the dorsal paraflocculus and the petrosal lobule of the monkey and in the dorsal paraflocculus and the rostral segment of the ventral paraflocculus of the cat. The C2 zone and a single D zone were distinguished in the paraflocculus (the petrosal lobule) of the rabbit cerebellum. The C2 zone, that extends over the entire cerebellum, occupies the lateral part of the flocculus in all species. Four zones 1-4 are present in the flocculus of the rabbit, three of these (1-3) are present in the primate flocculus. Of the 6 floccular climbing fiber (f) zones in the cat, fl corresponds to zone 4 of the rabbit, f2+3 to zone 3, f4 to zone 2 and f5+6 to zone 1. The zonal pattern of the flocculus proceeds for some distance on the paraflocculus. This transitional segment corresponds to folium p of the rabbit, to the ventral paraflocculus of the monkey and to the medial extension of the ventral paraflocculus (ME) of the cat. The lateral border of the hemisphere (heavy line) continues in the medial border of the flocculus. The inner surface of the cortex is shaded. Asterisks indicate the area without cortex in the center of the ansiform lobule. 1-4 = floccular zones 1-4 of rabbit and monkey; ANS = ansiform lobule; A N T anterior lobe; C 2 -~ zone C2; D1,2 = zones dl,2ME = medial extension of the ventral paraflocculus fl-6 = climbing fiber zones 1-6 of the flocculus of the cat - 9F I P L - posterolateral fissure; FLO - flocculus; FP folium P (rabbit); P F L D = dorsal paraflocculus; PFLV = ventral paraflocculus; P M D - paramedian lobule; POST - posterior lobe.

Two strips of Purkinje cells were distinguished by Umetani and Tabuchi (1988) in the vermis of lobule VIII (Fig. 153). The medial zone projected to both subdivisions of the fastigial nucleus, the second zone to the posterior interposed and lateral vestibular nuclei. This region, therefore, may include equivalents of the X and B zones of the anterior lobe. It was bordered on its lateral side by the C1 zone. The zonal organization of the lobules IX and X was analysed by Bernard (1987) and Tabuchi et al. (1989) in the rat. According to Bernard (1987) the zonal arrangement in lobule IX is very similar to the pattern in cat and rabbit with a medial zone projecting to the fastigial nucleus and a middle zone connected with medial portions of the interposed nuclei, corresponding with the posterior interposed nucleus. The most lateral zone 216

The cerebellum." chemoarchitecture and anatomy

Ch.I

Olivocerebettar projection Katayoma gNisimaru '88 BOtQLan g Henry '88 Tan et at.'94

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Fig. 150. Diagram of the afferent olivocerebellar projection according to Katayama and Nisimaru (1988) and the efferent projection of the zones of the nodulus to the vestibular and cerebellar nuclei according to Wylie et al. (1994) in the rabbit, fl = group fl of the medial accessory olive; DC = dorsal cap of Kooy; F = fastigial nucleus; IP = posterior interposed nucleus; MV = medial vestibular nucleus; P cell = Purkinje cell; SV = superior vestibular nucleus; VLO = ventrolateral outgrowth; I-VI = zones of rabbit nodulus, numbered according to Katayama and Nisimaru (1988). was connected with the ventral, parvicellular portion of the lateral cerebellar nucleus. The medial two zones shifted to a more lateral position in lobule X, where an additional zone, projecting to the medial vestibular nucleus, took up the most medial position. These zones also project in a differential manner, to the superior and descending vestibular nuclei and to the group y. The most lateral zone, that was connected with the lateral cerebellar nucleus, extended in lobule Xa and did not project to the vestibular nuclei. Although the Zebrin pattern in the caudal vermis is quite distinct and some correlations with the corticonuclear projection zones are obvious (Fig. 145), no precise comparisons with the Zebrin pattern have been made. Our knowledge of the cortico-vestibular and nuclear projection of the caudal vermis, therefore, is still deficient. Progress can be expected from anatomical and electrophysiological studies using the Zebrin pattern as a reference. 6.2. R E G I O N A L D I F F E R E N C E S IN T H E D E V E L O P M E N T O F T H E CEREBELLUM The development of the cerebellar cortex, the cerebellar nuclei and the precerebellar nuclei has been documented and reviewed in a series of papers on the rat by Altman (1975a,b,c) and Altman and Bayer (1985a,b, 1987a,b). Here we shall be concerned with 217

Ch. I

J. Voogd, D. Jaarsma and E. Marani

A

Flocculus

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Fig. 151. Diagram of the corticovestibular projections from flocculus, nodulus and uvula in Galago. Note complementarity between the projections of the flocculus and the caudal vermis, lvn = lateral vestibular nucleus; mvn = medial vestibular nucleus; spvn = spinal vestibular nucleus; svn= superior vestibular nucleus. Haines (1977a).

some aspects of regional differentiation in the development of the Purkinje cells of the cerebellar cortex. Several authors noticed that developing Purkinje cells are clustered into a number of parasagittal zones during early stages of cerebellar development, prior to the stage when the first fissures make their appearance. Purkinje cells and the cells of the cerebellar nuclei are generated in the ventricular layer (Jakob, 1928; Miale and Sidman, 1961), subsequently they migrate to the meningeal surface of the cerebellum where they settle in the cortical plate, deep to the external granular layer. Clustering of Purkinje cells in the cortical plate has been observed by a number of authors in mammals (Korneliussen, 1967, cetacea; Brown, 1985b; Brown et al., 1986, cat; Korneliussen, 1968b; Altman and Bayer, 1985b, rat; Hochstetter, 1929; Korneliussen, 1968c; Maat, 1978, 1981; Marani and Mai, 1992, man; Marani et al., 1986, rabbit; Kappel, 1981, monkey) and birds (Feirabend et al., 1976; Feirabend, 1983). Cell strands interconnect the Purkinje cell clusters with the cerebellar nuclei. The pattern that evolves from the position of the clusters in the cortical plate and their corticonuclear relations, is rather similar to the adult pattern of longitudinal corticonuclear projection zones (Korneliussen, 1967, 1968b; Kappel, 1981; Feirabend, 1983; Marani, 1986; Marani et al., 1986). The borders between the Purkinje cell clusters become indistinct when the fissures begin to develop and the 218

The cerebellum." chemoarchitecture and anatomy

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Fig. 152. Diagrams showing the topographic pattern of the projections from the various mediolateral levels of the tuber vermis (lobule VII) and the paramedian lobule to the cerebellar nuclear complex in the rat. A. Schematic diagram of the posterior surface of the cerebellum and subdivision of the tuber vermis and paramedian lobule, based on the topography of their projections. B. Schematic sagittal diagrams of the nuclear complex showing the terminal fields which receive projections from the individual subdivisions of the tuber vermis and paramedian lobule. AIN = anterior interposed nucleus; cm = caudomedial sub-division of the medial nucleus; Cop. pyr = copula pyramidis; D L H = dorsolateral hump; DLP = dorsolateral protuberance of the medial nucleus; LN - lateral cerebellar nucleus; LVN = lateral vestibular nucleus; m = medial nucleus; PIN = posterior interposed nucleus; Pml = paramedian lobule. Umetani (1989).

internal granular layer appears. Jacob (1928) and Hayashi (1924), were among the first to detect regional differences in corticogenesis in the human cerebellum (Fig. 154) and to correlate their corticogenetic zones with the corticonuclear projection to the different cerebellar nuclei (see Section 6.). A large time gap separates the stage when the borders between the clusters disappear, from the moment when the adult connections of the Purkinje cell have become established. Several methods have been employed to bridge this gap and to trace the developmental Purkinje cell patterns into adulthood. Altman and Bayer (1985b) in the rat, Brown (1985b) in the cat and Feirabend et al. (1985) in the chicken, using tritiated thymidine autoradiography, found evidence for differences in the time of birth between populations of Purkinje cells with a zonal distribution. According to Feirabend et al. (1985) some of the Purkinje cells of the lateral cerebellum and the cells of the lateral part of the central nuclei of the chicken are generated later than medially located cells. Zonal [3H]thymidine labelling of the Purkinje cells can be traced till after hatching. Several markers for adult Purkinje cells have been used to trace back their origin. Wassef and Sotelo (1984) studied the expression of cyclic GMP-dependent protein kinase (cGK) immunoreactivity during development of the rat (Fig. 155). They found a transient heterogeneity of the Purkinje cells for an antibody against cyclic GMP-dependent 219

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J. Voogd, D. Jaarsma and E. Marani Tuber vermis

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protein kinase (cGK) (De Camilli et al., 1984). Some clusters of Purkinje cells and their axons react with the antibody, while others are still negative. Some clusters could be recognized before their cells had completed their migration to the surface. Shortly after birth all Purkinje cells become cGK positive. Similar observations were made by Wassef et al. (1985) with two other Purkinje cell specific antibodies against calbindin-D28k (Legrand et al., 1983), and the Purkinje cell specific glycoprotein (PSG) (Langley et al., 1982) and by VanDaele et al. (1991) and Smeyne et al. (1991) for the transient zonal patterns in the development of the Purkinje cell-specific marker L7 (see Section 3.1.8.). Each antibody gave a different mosaic of positive and negative Purkinje cell clusters and this chemical heterogeneity disappeared shortly after birth, when all Purkinje cells became positive for the different markers. Some degree of heterogeneity is also present during the development of immunoreactivity for anti-parvalbumin in Purkinje cells of birds (Braun et al., 1986), but the early stages of development were not included in this study. Wassef et al. (1987) and Edwards et al. (1994) noticed that the surviving Purkinje cells in mice with mutations affecting their postnatal survival, are arranged in sagittal bands. This differential sensitivity of these Purkinje cells may be related to their chemical heterogeneity. Immunoreactivity for Zebrin (Hawkes and Leclerc, 1986) is not present before birth (Leclerc et al., 1988) and the reactivity for the enzyme 5'-nucleotidase, which is distributed in a similar zonal pattern in the molecular layer of the mouse also appears postnatally (Hess and Hess, 1986). The expression of the L7 gene in mouse Purkinje cells displays a similar develop220

The cerebellum: chemoarchitecture and anatomy

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mental heterogeneity. It first appears in paired bands in the vermis and the flocculus, later in clusters in the hemispheres. Ultimately it is present in all Purkinje cells (Smeyne et al. (1991)). Oberdick et al. (1993) slowed down this process by manipulation of the promotor region (Fig. 156). The transient expression of L7 in Purkinje cell bands with high and low levels of expression, as shown in whole mounts of the mouse cerebellum, is very similar if not identical to the Zebrin pattern as documented for adult mice by Eisenman and Hawkes (1993) (Fig. 139). It is not known whether the developmental patterns for other general Purkinje cell markers also resemble the Zebrin pattern. Moreover, it was demonstrated by Oberdick et al. (1993) that the L7 pattern also develops in vitro and, therefore, is independent of the ingrowth or the presence of extracerebellar afferents. Similar observations were made by Leclerc et al. (1988) for the development of the Zebrin pattern in rats, after section of the extracerebellar afferents. These results strongly advocate a primary role for the Purkinje cells in the development of zonal patterns (Wassef et al., 1992c). Few signs have been noticed of longitudinal subdivisions in the development or in the adult granular layer. The zonal distribution of AChE in the development of this layer was discussed by Marani (1986). Monoclonal antibodies against Stage Specific Embryonic Antigen-1 (SSEA-1), also known as the X-hapten, Lacto-N-Fucopentax III or FAL (Fucosyl-N-Acetyl Lactosamine), showed a longitudinal subdivision in the mitotic external granular layer of the rabbit cerebellum. This pattern was present from embryological day E8-E9 till postnatal day P15-16 and consisted of alternating positive and negative strips. The positive staining for SSEA-1 antibodies is exclusively present at the cell membrane of cells in the external granular layer (Marani and Tetteroo, 1983, Marani, 221

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Fig. 155. Staining with cyclic 3',5'-guanosine monophosphate dependent protein kinase (cGK) antisera of sections of cerebellum of rat fetuses of embryonic day E17, E19 and a neonate (PO) cut in the frontal plane. A,B. E 17. Cluster I is composed of a medial sheet (arrow in B) lying against the germinative neuroepithelium. Close to the midline this sheet bends dorsally and reaches the cortex. The central cluster (CC) is located at the center of the hemicerebellum. C. E 19. In this section four of the five cGK-positive clusters I-V are present. The labelled fiber-like material, which tails the labelled clusters (* and o) indicates the migration pathw.ays followed by the Purkinje cells of the clusters I and III from the subventricular plate and the central cluster at E17 to their present, superficial position. D. PO rat pup. Fiber bundles linking the clusters I and III with the cerebellar nuclei intersect at the former position of the central cluster. It is suggested that the bundle from cluster III (*) terminates in the dorsolateral protuberance. In the adult this connection corresponds to the projection of the lateral extension of the A zone of Buisseret-Delmas (1988a, compare Figs. 142 and 144). Bar in A - 100 ~m, in B, C and D = 500 ~tm. Wassef and Sotelo (1984). (

1986; Marani et al., 1986). These results were confirmed with several antibodies against SSEA-1. In dissociated rabbit cerebellum the only cell type that was found to express this antigen was the granule cell (Marani and Tetteroo, 1983, Marani et al., 1983, 1986). These results demonstrate the presence of a longitudinal pattern in an intrinsic mitotic

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Fig. 156. Effects of promotor gene truncation on L7-1acZ banding pattern in mice. Cerebella were dissected free from the rest of the brain and stained in whole mount. Cerebella are viewed from posterior (POST) and anterior (ANY). Cerebella were taken from postnatal day 11 animals carrying 4 kb (top row), 500 bp (middle row) and 350 bp (bottom row) promoter constructs. The patterns are very similar to the Zebrin pattern in mouse cerebellum (compare Fig.139). Expression of L7 is absent or low in P1 +, P3 +, P5 + (indicated with PIN in bottom panels) and P7 + in the 500 and 350 bp constructs. In P4+ there is a strong expression of L7 in the 500 bp construct, and a weak expression in the 350 bp construct. Reversed levels of expression are observed for the region of P4b + and P5a +. In the 500 bp construct they are weakly stained (but the P4b + and P5a + bands are visible as separate strips), in the 350 bp construct there is a strong expression of L7 over the entire area of these bands indicated with FN in lower pannel). Oberdick et al. (1993), interpretations by the authors of this chapter.

223

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Ch. I

Fig. 157. Configuration of the inferior olive and organization of the olivocerebellar projection in the cat. Olivary subnuclei and their projection areas in the cerebellum and the cerebellar nuclei (upper diagram) are indicated with the same symbols. The unfolded inferior olive is illustrated in the bottom diagram. 1-4 = in diagram of the cerebellum: lobules 1-4 of Bolk; in diagram of the inferior olive: projections to lobules 1-4;fl = group beta; D = dorsal accessory olive; d.cap = dorsal cap; d.1. = dorsal leaf of the principal olive; dm.c.col. = dorsomedial cell column; 1= lateral; lob a = lobulus a of Bolk (X of Larsell); lob b = lobulus b of Bolk (IX of Larsell); lob c l = lobulus c l of Bolk (VIII of Larsell); lob c = lobulus c of Bolk (VI + VII of Larsell); lob.simpl. - lobulus simplex; M = medial accessory olive; m = medial; N.fast = fastigial nucleus; N.int = interposed nucleus; N.lat = lateral cerebellar nucleus; v.1. = ventral leaf of the principal olive; v.l.o. = ventrolateral outgrowth. Brodal (1940). (

layer of the cerebellum which may be related to the establishment of transient contacts between ingrowing afferent fibers or Purkinje cell dendrites and external granule cells. 6.3. T H E O R G A N I Z A T I O N O F T H E O L I V O C E R E B E L L A R P R O J E C T I O N The olivocerebellar projection has been studied mainly in the cat (see Brodal and Kawamura, 1980 and Voogd, 1982 for reviews). Fewer data are available for primates and sub-primates (Brodal and Brodal, 1981, 1982; Whitworth et al., 1983; Whitworth and Haines, 1986b). A more complete picture of the olivocerebellar projection in the rat is emerging (Buisseret-Delmas and Angaut, 1993; Ruigrok and Cella, 1995; Voogd, 1995). The evidence on the role of excitatory aminoacids as the transmitter in the olivocerebellar pathway was considered in Section 3.2.2. It was concluded that glutamate is the most likely transmitter of the climbing fibers. The configuration and ultrastructure of the inferior olive, the organization of the afferent connections of the olive, the olivocerebellar projection and the topographical distribution of peptidergic climbing fibers will be reviewed in the next sections.

6.3.1. Configuration and ultrastructure of the inferior olive The morphology and the subdivision of the inferior olive have been described in the comparative anatomical studies of Kooy (1917), Mar6schal (1934) and Whitworth and Haines (1986a). Brodal's (1940) subdivision of the inferior olive in the cat and his mode of representation of the olive as imagined unfolded in one plane (Fig. 157) have become generally accepted. Dorsal accessory (DAO), medial accessory (MAO) and principal (PO) subnuclei can be distinguished in most mammalian species (Figs 158 and 159). The dorsomedial subdivision of the caudal M A O was indicated as group beta by Brodal (1940). Three parallel, longitudinal cell columns, indicated from laterally to medially as a, b and c (c is the equivalent to the group beta) were distinguished in the caudal M A O of the macaque monkey (Bowman and Sladek, 1973). F o u r columns (a, b, c and beta) were delineated in the caudal M A O of the rat (Gwyn et al., 1977). This apparent discrepancy was solved by Frankfurter et al. (1977) and Ikeda et al. (1989), who showed that subnucleus b in squirrel and macaque monkeys can be subdivided into medial and lateral parts and that only the medialmost region of this subnucleus is connected with the oculomotor vermis (lobule VII) and, therefore, corresponds with subnucleus c in the rat (Hess, 1982b; Akaike, 1992). The rostral half of the M A O sometimes is indicated as the rostral extension of subnucleus a, but this does not serve a useful purpose, because its connections differ 225

Ch. I

J. Voogd, D. Jaarsma and E. Marani

substantially from the caudal half of the MAO. The dorsomedial cell column (DMCC) of the cat is located dorsomedial to the rostral half of the medial accessory olive. Bowman and Sladek (1973) identified their cell group g, which is connected to the medial tip of the ventral leaf of the PO, as the DMCC in the macaque monkey. The DMCC of the rat has been identified as a similar group, connected with the ventral leaf of the PO (Gwyn et al., 1977). However, since the DMCC is part of the MAO and forms a rostral continuation of the group beta in most species (Whitworth and Haines, 1986a),

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Diagrams of the inferior olivary complex of the rhesus monkey. Dorsal accessory olive is shown in black. Medial accessory olive and attached groups (nucleus fl and dorsomedial cell column) are hatched. Principle olive, dorsal cap, and ventrolateral outgrowth are left white. Subnuclei a, b, and c of the medial accessory olive according to the nomenclature of Bowman and Sladek (1973). A series of drawings of 13 equally spaced transverse sections through the olive in the upper panel is represented from rostrally (I) to caudally (XIII). In the lower panel the inferior olive is represented as imagined unfolded (see diagram at bottom of the Figure). In the drawings of the sections showing the principal olive, borders (broken lines) are indicated between the dorsal, lateral, and ventral lamella in accordance with Bowman and Sladek (1973). The solid line at the lateral convexity of the principal olive corresponds to the vertical line in the diagram of the unfolded olive, and is used for orientation. In this diagram broken lines in the medial accessory olive refer to borders betwcen regions which are not clearly separated. In the principal olive broken lines indicate the arbitrary separations between dorsal, lateral and ventral lamella shown in the drawings of transverse sections. The dorsomedial cell column is hidden behind the medial accessory olive (dotted outlines). Brodal and Brodal (1981). Fig. 159.

227

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J. Voogd, D. Jaarsma and E. Marani

another cell group positioned directly dorsomedial to the MAO, has been identified as the DMCC in the rat (Azizi and Woodward, 1987; Bernard, 1987; Ruigrok and Voogd, 1990). Rostrally the DMCC of both sides seem to merge. The cell group connected to rostral leaf of the PO of the rat was indicated as the dorsomedial group (DM) (Fig. 158). A dorsal and a ventral leaf usually are distinguished in the PO: in the cat the ventral leaf is continuous with the medial pole of the dorsal accessory olive. In other mammals the DAO is continuous with the dorsal leaf and the ventral leaf ends as the DMCC in macaque monkeys and at the DM in rats. The ventral leaf of the PO caudally tapers into the ventrolateral outgrowth (VLO) that continues as the dorsal cap (DC) of Kooy (1917), located dorsal to the group beta. In dorsal view the DAO of the inferior olive of the cat and the monkey is boomerangshaped, with a narrow, medially directed, caudal tail. In the rat the caudal pole of the DAO consists of a dorsal and a ventral fold, that are continuous medially. The dorsal fold is only present at caudal levels, the ventral fold extends to the rostral tip of the DAO (Azizi and Woodward, 1987) (Fig. 158). Ovoid perikarya in Nissl-stained sections of the inferior olive of the cat are dispersed in clusters comprising up to eight neurons (Sotelo et al., 1974). The dendrites of typical Golgi-impregnated olivary neurons recurve towards the cell body (Fig. 160). This type of neuron, with a compact dendritic arbor, is the main cell type of the principal olive and the rostral pole of the MAO. Other subdivisions of the inferior olive contain a mixture of compact neurons and neurons with long, unramified dendrites. The latter type of neuron was mainly found in the caudal MAO, group beta and the DC (Scheibel and Scheibel, 1955; Scheibel et al., 1956; Ruigrok et al., 1990). Dendrites of both types of neurons give rise to long and thin branching spines (Bowman and King, 1973; Sotelo et al., 1974; Gwyn et al., 1977). The ultrastructure of the inferior olive was reviewed by De Zeeuw (1990) (Fig. 161). Glomeruli are characteristic features of the olivary neuropil (Nemecek and Wolff, 1969; Bowman and King, 1973; Sotelo et al., 1974; King et al. 1975; King, 1976; Gwyn et al., 1977; Rutherford and Gwyn, 1980; Bozhilova and Ovtscharoff, 1979). Glomeruli contain a core of dendritic spines contacted by axon terminals and surrounded by a glial capsule. On average, the glomeruli contain spines derived from six different neurons. Sometimes spines of the initial axonal segment are incorporated together with the dendritic spines in the same glomerulus (De Zeeuw et al., 1990a,b). Small gap junctions, with a heptalaminar structure including a 2 nm wide cellular space, were observed between spines in and outside glomeruli (Sotelo et al., 1974, 1986). De Zeeuw et al. (1990a) excluded the presence of gap junctions between elements of the same neuron. The importance of dendrodendritic coupling is further supported by the presence of dendritic lamellar bodies that can be associated with dendrodendritic gap junctions. These organelles are ubiquitously distributed in all olivary subdivisions and their density is higher in the olive than in any other brain area (De Zeeuw et al., 1995). Another form of aggregation of dendrites of olivary neurons is the dendritic thicket (Sotelo et al., 1974; 1986, Molinari, 1987; De Zeeuw et al., 1993). Thickets are formed by several dendrites in direct apposition with each other, but without any dendrodendritic membrane specializations. The extent of the electrotonic coupling of olivary neurons was studied by injecting lucifer yellow into single cells in slices of the guinea pig brain stem. Transfer of lucifer yellow via the gap junctions between dendrites, resulted in the labelling of aggregates consisting of up to 5 cells (Bernardo and Foster, 1986). The maximal spatial extent of electrotonic coupling in the inferior olive has not been determined, but may extend 228

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Fig. 161. GABAergic terminals derived from the cerebellar nuclei innervate dendritic spines (asterisks) in the glomeruli of the principal olive and medial accessory olive in the cat. The double labelled terminals can be recognized by the 15 nm gold particles (GABA) and the reaction products of the anterogradely transported WGA-HRP (large arrows). Note in B and C that cerebellar GABAergic terminals are directly apposed to dendritic spines coupled by gap junctions (small arrows). Open arrows indicate symmetric synapses. Scale bar in A = 0.31 r in B = 0.18 r in C = 0.35 r De Zeeuw et al. (1989b).

230

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beyond the classical morphological borders of the neuropil of the subdivisions of the inferior olive. De Zeeuw et al. (not yet published) demonstrated that synchrony between olivary neurons that project to the left and right Crus IIa of the rat in part can be explained by direct coupling through cells in the DMCC that bridge the left and right rostral DAO. Synchronization of activity in olivary neurons derived from different subdivisions is also supported by studies from Sj61und et al. (1980) and Lang et al. (1995) who induced a hypersynchronization of the climbing fiber discharge in the projections of particular olivary subnuclei by the administration of harmalin. Three types of axon terminals were recognized in the neuropil of the olive. It was found in aldehyde-fixed material postfixed in osmium that about half of the terminals contained spherical vesicles and were associated with asymmetrical synapses (Bowman and King, 1973; Mizuno et al., 1974; Sotelo et al., 1974; Gwyn et al., 1977; Rutherford and Gwyn, 1980). A second type contains pleomorphic vesicles and is associated with symmetrical synapses. A third type of terminal contains dense core vesicles. Most of the terminals that can be double labelled with an antegrade tracer from the cerebellar nuclei and an antibody against a conjugate of GABA belong to the second type, with pleomorphic vesicles and asymmetrical synapses (Angaut and Sotelo, 1989; De Zeeuw et al., 1988, 1989a and b, 1992, 1994b, Fig. 161). At present the cerebellar and vestibular nuclei, the nucleus prepositus hypoglossi, the parasolitary nucleus and the cuneate nucleus are the only known sources for this type of GABAergic terminal, and most of these terminals disappear after lesions of these nuclei (Nelson and Mugnaini, 1985; Nelson et al., 1986; Fredette and Mugnaini, 1991; De Zeeuw et al., 1993). Less frequently, GABAergic terminals with so-called crest-synapses can be observed (De Zeeuw et al., 1994b). These terminals occur relatively often in the DC and VLO and are derived from the area of group y and the ventral dentate nucleus (Fig. 162). Two other populations of GABA-containing terminals exist. One minor population contains clear, oval vesicles and sometimes was found apposed to the perikaryon. It was infrequently labelled from the cerebellum (De Zeeuw et al., 1989b). GABA-like immunoreactivity was also found in certain terminals of the granular type, that contained a large number of dense core vesicles. The granular terminals rarely made synaptic contacts. They could not be labelled from the cerebellum and they were always found outside the glomeruli, sometimes apposed to somata. The same type of granular terminal was labelled in the inferior olive with [3H]serotonin (Wiklund et al., 1981a) or antibodies to conjugates of serotonin (King et al., 1984). It seems likely, therefore, that serotonin and GABA are co-localized in a subpopulation of these terminals. All other reported non-cerebellar afferents terminate with boutons containing spherical vesicles (nuclei at the mesodiencephalic junction: King et al., 1978; Cintas et al., 1980; De Zeeuw et al., 1988, 1989a,b; spinal cord: King et al. 1976; Mizuno et al., 1976; Gwyn et al., 1983; Molinari and Starr, 1989; Molinari, 1988). Most of these terminals were apposed to distal dendrites or spines inside and outside glomeruli, relatively few were found to contact somata. Afferents from the spinal cord and the gracile nucleus never have been found in glomeruli in close contact with gap junctions (Molinari and Starr, 1989; Molinari, 1987, 1988; Molinari et al., 1990). The preferential axo-dendritic mode of termination of dorsal column afferents may be due to the relative paucity of spines on the long, radiating dendrites of the cell type that prevails in the spinal areas of the inferior olive. Axon collaterals from olivary neurons that terminate in the same and the contralateral inferior olive have only been observed in young kittens (Ramon y Cajal, 1911) and in cases of olivary hypertrophy in the cat (Ruigrok et al., 1990; De Zeeuw et al., 1990d). 231

Ch.I

J. Voogd, D. Jaarsma and E. Marani

Fig. 162. Double labelled terminals (GABA and anterogradely transported WGA-HRP) from the contralateral PrH terminate on distal and proximal portions of the neurons in the dorsal cap of the rat (A and B) and the rabbit (C). In A, a double labelled terminal (right) and a GABAergic large granular terminal (left) are apposed to dendritic spines (asterisks) coupled by a gap junction (small arrows). In B and C the terminals are apposed to somata. The large arrows indicate the WGA-HRP reaction products. The open arrows and the arrowhead indicate symmetric synapses and an asymmetric synapse, respectively. Scale bar in A - 0.24 r in B = 0.41 r in C = 0.39 r De Zeeuw et al. (1994).

Cerebellar, GABAergic and mesodiencephalic, non-GABAergic terminals contacted both spines inside glomeruli and dendritic shafts. The cerebellar (King et al., 1976; Angaut and Sotelo, 1987) and/or GAD or GABA-immunoreactive (Sotelo et al., 1986; Fredette and Mugnaini, 1991) terminals are associated with gap junctions inside the glomeruli. De Zeeuw et al. (1989a,b; 1990a and c) concluded that both GABAergic and 232

The cerebellum." chemoarchitecture and anatomy

Ch. I

non-GABAergic mesodiencephalic terminals in the PO and the rostral MAO of cat and rat innervate the same spines including those that are electrotonically coupled. However, the cerebellar GABAergic terminals, but not the mesodiencephalic terminals, showed a strong preference to be strategically located next to both dendrites coupled by a gap junction (Fig. 161). Most or all of the terminals contacting the somata of olivary neurons contained pleomorphic vesicles (Bowman and King, 1973; Gwyn et al., 1977; Rutherford and Gwyn, 1980; Sotelo et al., 1986). Moreover, large GABAergic granular terminals were observed next to the soma. The proportion of somatic GABAergic terminals, that could be labelled from the cerebellar nuclei was significantly lower than among the terminals in the entire neuropil (De Zeeuw et al., 1989b). The presence of a non-cerebellar, GABAergic innervation of the cerebellum was also suggested by Nelson and Mugnaini (1985) and Fredette and Mugnaini (1991), who showed that total cerebellectomy does not cause a total depletion of GAD-positive terminals in the olive of the rat, and by the electrophysiological studies of Andersson et al. (1988) and Weiss et al. (1990). The GABAergic innervation of the axon hillock appears to be of cerebellar origin (De Zeeuw et al., 1990b). Possible extracerebellar sources for a GABAergic innervation of the inferior olive include the raphe nuclei and the adjacent reticular formation (Bishop, 1984), the nucleus parasolitarius and the cuneate nucleus (Nelson and Mugnaini, 1989) and the intrinsic GABAergic neurons of the inferior olive. Intrinsic GABAergic neurons of the inferior olive seem to be rare. Mugnaini and Oertel (1985), Nelson and Mugnaini (1989) and Fredette et al. (1992) found only few GAD-positive cells in the inferior olive of the rat. These cells were usually small and provided with unbranched, ramifying dendrites. They were differently distributed in different species; in the cat most occur in the rostral tip of the MAO and in the dorsal fold of the DAO, in primates in the PO (Fredette et al., 1992). They were more numerous in the baboon, where they accounted for 5% of the neurons of the inferior olive (Walberg and Ottersen, 1989). Colchicine injections in rat, rabbit and cat enhanced their visibility and revealed a high density of GAD-positive neurons in the reticular formation directly dorsal to the inferior olive (Nelson and Mugnaini, 1989; Fredette et al., 1992). Large cells with dendrites penetrating in the neuropil of the olive, which resemble the cells of the reticular formation, were identified by Sotelo et al. (1974) in the cat. Similar, peri-olivary cells, containing glutaminase were illustrated by Kaneko et al. (1989) in the rat. Bishop (1984) and Bishop and King (1986) demonstrated that dendrites of intracellularly injected reticular neurons can contribute to the olivary glomeruli. 6.3.2. Afferent connections of the inferior olive

Afferent systems of the inferior olive have been reviewed by Brodal and Kawamura (1980) for the cat and by Martin et al. (1980) for the opossum. For the rat the tabulated summary of its afferent connections in the paper of Brown et al. (1977) and the review by Flumerfelt and Hryccyshyn (1985) are useful. Afferent systems of the inferior olive can be subdivided into three groups: (1) the GABAergic nucleo-olivary and vestibuloolivary projections; (2) the monoaminergic and cholinergic projections to the inferior olive; (3) the specific projections from the spinal cord, certain brain stem nuclei and the cerebral cortex will not be considered in this chapter. Their neurotransmitters are not known.

233

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6.3.2.1. The nucleo-olivary and vestibulo-olivary projections." The GABAergic afferents of the inferior olive The origin of the nucleo-olivary projection from the cerebellar nuclei (Section 5.2.) and their mode of termination (Section 6.3.1 .) has been considered before. In this section the topographical organization of the nucleo-olivary and the closely allied vestibulo-olivary projections will be reviewed. The nucleo-olivary projection is entirely GABAergic, the vestibulocerebellar projection only partially so. For the main part the nucleo-olivary and the olivocerebellar projections are reciprocally organized, in the sense that the cerebellar nuclei which are the target of a particular olivo-corticonuclear loop give rise to a nucleo-olivary projection to this particular olivary subnucleus. This reciprocity is not complete, because the olivary projections of some of the cerebellar nuclei terminate bilaterally in the olive. The olivary projection from Deiters' nucleus clearly reciprocates the olivocerebellar projection to the B zone, but for the vestibulo-olivary projections from other vestibular nuclei this reciprocity is less obvious. Nucleo-olivary fibers terminate in all parts of the inferior olive. Roughly the density of the GABAergic boutons in different subnuclei of the olive represents the density of the nucleo-olivary projection to these parts. The density of GAD immuno-reactivity is higher in the group beta of the rat, which also contains the largest reactive boutons, followed by the subnucleus c of the MAO, the DMCC, the medial and lateral poles of the DAO, the bend of the PO, the DC and the DM (Sotelo et al., 1986) (Fig. 166). Nelson and Mugnaini (1988) noticed that the dorsal fold of the rat DAO contains a higher immunoreactivity than the ventral fold and confirmed the difference in bouton-size between strongly immunoreactive and less reactive regions. Nelson et al. (1989) found the same distribution in a comparative study of GAD-immunoreactivity of the olive in rabbit, cat, rhesus monkey and man (Fig. 163). Activity is highest in the beta nucleus. The DAO contains several, differently stained regions, with high immunoreactivity in the rostromedial, and in the caudolateral parts of the nucleus. Staining in the caudal MAO is lowest in the central subnucleus b and higher in the lateral subnucleus a and the rostral portion of the MAO. Immunoreactivity of boutons in the DC (high in rabbit and monkey; low in rabbit and man) and the DMCC (high in rabbit and cat, and low in monkey) differs for different species. GAD-immunoreactive boutons disappear from the contralateral PO, the rostral MAO and the lateral half of the ventral fold of the DAO of the rat after chronic lesions of the cerebellar nuclei or the superior cerebellar peduncle in the rat. The dorsal fold of the DAO is depleted of GAD-positive boutons after lesions extending into the lateral vestibular nucleus (Fredette and Mugnaini, 1991). Additional destruction of the vestibular nuclei results in the disappearance of GAD from the group beta but not from the medial half of the ventral fold of the DAO and the caudal MAO (Nelson and Mugnaini, 1989). GAD-immunoreactive neurons in the parasolitary and cuneate nuclei could be labelled after injection of retrograde tracers in the inferior olive of the rat (Nelson and Mugnaini, 1989). These nuclei, therefore, provide additional GABAergic projections to the inferior olive. The projection of the ipsilateral parasolitary nucleus was located in the medial subnucleus c of the caudal MAO by these authors. Connections from the parasolitary nucleus (indicated as the lateral solitary nucleus) also were documented in earlier studies by Loewy and Burton (1978) and Molinari (1985) in the cat. The inhibitory connections from the cuneate nucleus in the rat are crossed and terminate in the medial DAO. A similar GABAergic cuneo-olivary pathway appears to be responsible 234

The cerebellum." chemoarchitecture and anatomy

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ble~ ~cies po~--DAO rostral Fig. 163. Schematic generalized diagram of GAD immunostaining intensities in regions of the mammalian IO based upon studies in rabbit, cat and Rhesus monkey. Intensity was visually graded on a scale of 5-1, where 5 indicates the most intense staining. The beta nucleus and medial aspect of the rostral DAO have the highest intensity in the IO (black regions). Most of the IO is stained with intermediate intensities (hatched regions). Subnucleus b of the MAO (white region) is least intensely stained. Staining in the dorsal cap and the dorsomedial cell column (dotted regions) varied by species, aMAO = subnucleus a of the medial accessory olive; beta = subnuleus beta; bMAO = subnucleus b of the medial accessory olive; DAO = dorsal accessory olive; dc = dorsal cap; dmcc = dorsomedial cell column; PO = principal olive; vlo = ventrolateral outgrowth. Nelson et al. (1989).

for the i n h i b i t i o n o f n e u r o n s in the D A O on s t i m u l a t i o n o f the red n u c l e u s or the c o n t r a l a t e r a l r u b r o s p i n a l t r a c t in the cat (Weiss et al., 1990). T h e t o p o g r a p h y in the c e r e b e l l a r n u c l e o - o l i v a r y p r o j e c t i o n has b e e n s t u d i e d in different m a m m a l i a n species. T h e n u c l e o - o l i v a r y fibers f r o m the i n t e r p o s e d a n d l a t e r a l nuclei r u n in a s e p a r a t e tract, v e n t r a l to the b r a c h i u m c o n j u n c t i v u m in cat ( L e g e n d r e a n d C o u r v i l l e , 1987), rat ( C h o l l e y et al., 1989) a n d r a b b i t (Tan et al., 1995b). F a s t i g i o - o l i v a r y 235

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J. Voogd, D. Jaarsma and E. Marani

and vestibulo-olivary fibers take other routes (Nelson and Mugnaini, 1989; Ruigrok and Voogd, 1990). It is generally assumed that the anterior- and posterior interposed nuclei project to the rostral DAO and MAO respectively and the lateral, dentate nucleus to the PO (cat: Tolbert et al. 1976b; Buisseret-Delmas and Batini, 1977; Dietrichs and Walberg, 1981, 1985, 1986; Courville et al., 1983a; monkey: Kalil, 1979; Asanuma et al., 1983; Gonzalo-Ruiz and Leichnetz, 1990; opossum: Martin et al., 1976, 1980; rat: Brown et al., 1977; Angaut and Cicirata, 1982; Haroian, 1982; Swenson and Castro, 1983 a and b; Nelson and Mugnaini, 1989; Ruigrok and Voogd, 1990). Points of discussion were, and still are the presence and the extent of the fastigio-olivary projection, the nucleo-olivary connections of subdivisions of the dentate nucleus and of certain cerebellar subnuclei in rat (dorsolateral protuberance of the fastigial nucleus, dorsolateral hump), the presence of ipsilateral nucleo-olivary projections and the relations of the cerebellar nucleo-olivary projection with the GABAergic and non-GABAergic vestibulo-olivary connections. A projection of the medial cerebellar nucleus to the caudal MAO was denied by Brown et al. (1977) and Haroian (1982) for the rat, but was found by Achenbach and Goodman (1968), Angaut and Cicerata (1982) and Swenson and Castro (1983a and b). In the cat negative findings were published by Graybiel et al. (1973), Tolbert et al., (1976b) and Courville et al. (1983a), but the connection was found to be present by

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236

The cerebellum." chemoarchitecture and anatomy

Ch. I

Sugimoto et al. (1980) and Dietrichs and Walberg (1985). In the monkey negative results were obtained in the anterograde tracer studies from the fastigial nucleus by Batton et al. (1977), Kalil (1979) and Asanuma et al. (1983). Ikeda et al. (1989), however, reported projections from the caudal part of the fastigial nucleus to medial portions of the caudal MAO in the macaque monkey. Ruigrok and Voogd (1990) in a recent study, using anterograde transport of Phasaeolus vulgaris lectin injected in the cerebellar nuclei of the rat, established projections from rostral regions of the fastigial nucleus to a lateral zone in the caudal MAO and of the caudal fastigial nucleus to the group beta (Fig. 164). The projection of the dorsolateral hump is located in a circumscribed area in the medial subnucleus c, that coincides with the tectal recipient zone of Akaike et al. (1992) that projects to the vermis of lobule VII and the lateral extension of the A zone in the hemisphere of the lobules VI and VII. The central portion of the caudal MAO (subnucleus b) lacks a nucleo-olivary projection. No terminations were found in the DC, which has been reported to receive fastigial projections by Angaut and Cicerata (1982), Swenson and Castro (1983a and b) in the rat and Dietrichs and Walberg (1985) in the cat. The results of experiments on nucleo-olivary projections from different parts of the dentate nucleus to the PO are conflicting. According to Beitz (1976) and Tolbert et al. (1976b) in the cat and Kalil (1979) in the monkey the dorso-ventral and medio-lateral relations are maintained in the nucleo-olivary projection of the dentate nucleus. Dietrichs and Walberg (1985) denied this and proposed a complicated rostro-caudal relationship between both structures. According to Angaut and Cicirata (1982) the medial dentate of the rat, including the dorsolateral hump, projects to the ventral leaf, and the lateral dentate to the dorsal leaf of the PO. Swenson and Castro (1983a and b) described a projection of the rostral dentate of the rat to the dorsal leaf and of the caudal dentate to the ventral leaf of the PO. Chan-Palay (1977) claimed that the projection in the monkey is reversed: caudal dentate projecting to the ventral PO. Some of her experiments, however, show an exclusive projection of the caudal dentate to the ventral leaf of the PO. Ruigrok and Voogd (1990) confirmed Angaut and Cicerata's (1982) topology for the rat (Fig. 164). They located the nucleo-olivary projection of the dorsolateral hump in the dorsomedial group (DM), the enlarged medial portion of the ventral leaf of the PO. The DMCC that was often confused with the DM of the rat, did not receive a nucleo-olivary projection. The nucleo-olivary projection is not completely crossed. Some fibers from the dorsolateral hump, IP and the ventromedial lateral nucleus recross at the level of the inferior olive, and terminate in the ipsilateral DM, rostral MAO and ventral leaf of the PO respectively. Ipsilateral labelling was sparse or absent in other parts of the inferior olive. Projections from the vestibular nuclei terminate in subnuclei that do not receive a nucleo-olivary projection, such as the DMCC, the group beta and the central region (subnucleus b) of the caudal MAO in cat (Fig. 165) (Saint-Cyr and Courville, 1979; Gerrits et al., 1985a), opossum (Martin et al., 1980) and rat (Nelson and Mugnaini, 1989). These projections are mainly crossed, and, in the rat at least, entirely or partially GABAergic. An ipsilateral projection from the superior vestibular nucleus to a lateral zone in the caudal MAO and the bend region of the PO in the cat was reported by Gerrits et al. (1985a). The uncrossed, GABAergic projection from the parasolitary nucleus to subnucleus c (Nelson and Mugnaini, 1989), and the bilateral projections from the nucleus prepositus hypoglossi to DC and VLO (Gerrits et al., 1985a; McCrea and Baker, 1985; De Zeeuw et al., 1993) should be included in the vestibulo-olivary projection. Terminations from the nucleus prepositus hypoglossi were located both in subnuclei dominated by descend237

Ch. I

J. Voogd, D. Jaarsma and E. Marani

PO

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Fig. 165. The vestibulo-olivary projection in the cat. Localization of terminal fields with antegrade axonal transport of [3H]leucine. A. Contralateral projection from the superior vestibular nucleus. B. Ipsilateral projection from the medial vestibular nucleus. C. Bilateral projections from the nucleus prepositus hypoglossi. MAO = medial accessory olive; PO = principal olive. Gerrits et al. (1985a).

ing visual afferent connections (DC and VLO) and in the caudal MAO, where they overlap with vestibular afferents from the medial and the spinal vestibular nuclei. The prepositus hypoglossi nuclear projection to the DC in rat and rabbit is bilateral and deals mainly with the caudal DC. It contains GABAergic (De Zeeuw et al., 1993) and cholinergic (Barmack et al., 1991) components. Recently it was shown that these two neurotransmitters are co-localized in the same terminals (De Zeeuw et al, unpublished). The GABAergic projections from the nucleus prepositus hypoglossi to the VLO and 238

The cerebellum." chemoarchitecture and anatomy

Ch. I

subnucleus c are exclusively contralateral and much sparser. The VLO and the rostral DC receive their major GABAergic input from the ventral dentate nucleus and the adjacent group y in the rabbit (De Zeeuw et al., 1994b). 6.3.2.2. Monoaminergic and cholinergic projections A plexus of varicose catecholaminergic fibers has been mapped in the inferior olive of cat, monkey and in several rodents with histofluorescence methods (Fuxe, 1965a and b; Hoffman and Sladek, 1973; Sladek and Bowman, 1975; Sladek and Hoffman, 1980). The distribution of these varicose fibers is heterogeneous and displays marked species variations. The distribution of serotoninergic fibers is also heterogeneous and speciesdependent, but it differs from the distribution of the catecholamines (Wiklund et al., 1977, 1981a,b; Sladek and Hoffman, 1980; Takeuchi and Sano, 1983; Bishop and Ho, 1984; King et al., 1984; Par6 et al., 1987; Compoint and Buisseret-Delmas, 1988). Immunocytochemical methods, using antibodies against conjugates of serotonin, were applied in the investigations reported in the last five papers. The catecholaminergic innervation of the inferior olive of the rat is strongest for the dorsal leaf of the PO and the rostral MAO (Fig. 166) (Sladek and Bowman, 1975). For serotonin the most intensely innervated subnucleus of the inferior olive of the rat is the lateral portion of the rostral lamella of the DAO (Fuxe, 1965a,b; Takeuchi and Sano, 1983; Bishop and Ho, 1984; Par6 et al., 1987; Compoint and Buisseret-Delmas, 1988), the dorsal leaf of the DAO is spared (Fig. 166) (Takeuchi and Sano, 1983). For the MAO and the PO the descriptions differ. Takeuchi and Sano (1983) found a somewhat higher innervation of the rostral MAO. Bishop and Ho (1984) described a dense innervation by varicosities of the subnuclei a and b of the caudal MAO, moderate numbers of serotonin-like immunoreactive fibers in subnucleus c and the caudal PO and few elements in group beta and the DC (Fig. 166). Par6 et al. (1987) confirmed this distribution for the caudal MAO. Compoint and Buisseret-Delmas (1988) stressed the presence of an immunoreactive plexus around the caudal MAO, with few fibers penetrating the center of the nucleus. Both catecholaminergic and serotoninergic fibers predominate in the lateral part of the caudal MAO of the cat. At more rostral levels only the medial MAO and the DMCC are densely innervated by serotoninergic fibers. The catecholaminergic and serotoninergic innervation of the DAO are complementary, with a serotoninergic plexus occupying the lateral and caudal DAO with the exception of its most lateral border region (Wiklund et al., 1977; Takeuchi and Sano, 1983). A central column of the caudal MAO contains a dense plexus of serotoninergic fibers in monkey (Takeuchi and Sano, 1983) (Fig. 167) and opossum (King et al., 1984). Other parts of the MAO of the opossum, including subnucleus c and the rostral pole of the MAO are less densely innervated. Serotoninergic fibers also predominate in the lateral DAO of the monkey (Takeuchi and Sano, 1983) but a serotoninergic plexus is present in the entire DAO of the opossum (King et al., 1984). The distribution of serotoninergic fibers in the PO differs for the different species, with an innervation of the ventral leaf of the cat (Sladek and Bowman, 1975), both the ventral and the dorsal leaf in the monkey (Takeuchi and Sano, 1983) (Fig. 168), and the dorsal leaf of the PO in the opossum (King et al., 1984). Serotonin was localized in axons and their terminals at the ultrastructural level by Wiklund et al. (198 l a) in the DAO of the rat with high resolution autoradiography of [3H]serotonin and by King et al. (1984) and Compoint and Buisseret-Delmas (1988) with 239

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Fig. 166. The innervation of the inferior olive of the rat by catecholaminergic, serotoninergic, substance P, and gamma-aminobutyric acid decarboxylase (GAD)-immunoreactive fibers and the distribution of acetylcholinesterase (ACHE). a = subnucleus a of the medial accessory olive; b = subnucleus b of the medial accessory olive; beta = subnucleus beta; D A O = dorsal accessory olive; d c = dorsal cap; dl = dorsal eaf principal olive; D M = dorsomedial subnucleus; dmcc = dorsomedial cell column; MAO = medial accessory olive; PO = principal olive; vl = ventral leaf principal olive; vlo = ventrolateral outgrowth; XII = hypoglossal nerve.

preembedding immunocytochemistry in the same species. The label was located in thin axons and varicosities, containing large dense core vesicles and small, clear vesicles or tubulo-vesicular elements. Few terminals engage in synaptic contacts (Wiklund et al.: 5%; King et al.: 2%), mainly with dendritic shafts, and never with the spines located in the glomeruli. Configurations suggesting axo-somatic contacts were found in the subnucleus b of the caudal MAO in the opossum (King et al., 1984). Similar terminals in the rostral MAO of the cat, containing large dark core vesicles, displayed GABA-like immunoreactivity (De Zeeuw et al., 1989b). The possibility that certain, non-cerebellar GABAergic projections are derived from the raphe nuclei or the adjoining medial reticular formation (Bishop 1984), and that they are co-localized with serotonin still needs to be explored. According to Compoint and Buisseret-Delmas (1988) the serotoninergic innervation of the inferior olive of the rat, at least in part, is derived from cell groups surrounding it. The nucleus raphe obscurus and the reticular formation dorsal to the olive innervate the MAO, the reticular formation and cell groups lateral to the DAO innervate the latter subnucleus. Co-localization of serotonin and substance P, that display a similar distribution in the olive of the rat

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Fig. 167. Schematic representation of the distribution of serotonin fibers in the inferior olivary complex of the monkey (a, rostral; i, caudal). Abbreviations: see Fig. 168. Takeuchi and Sano (1983).

241

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Fig. 168. Schematic representation of the distribution of serotonin fibers in the inferior olivary complex of the cat (a, rostral; i, caudal). B = group beta; DA--DMC = dorsomedial cell column; DAO = dorsal accessory olive; DC = dorsal cap; MAO = medial accessory olive; PO = principal olive; POD - dorsal lamella of the principal olive; POV = ventral lamella of the principal olive. Takeuchi and Sano (1983).

(Bishop and Ho, 1984; Par6 et al., 1987) (Fig. 166), is another possibility (H6kfelt et al., 1978; Pelletier et al., 1981; Par6 et al., 1987). A differential distribution of AChE has been described in the neuropil of the inferior olive of cat (Marani et al., 1977) (Fig. 169), ferret, rabbit (Marani, 1986,), rat (Fig. 166), and the opossum (Martin et al., 1975). These distributions are fairly consistent as to the columnar distribution of the enzyme in the caudal MAO and the rostral DAO, the absence of AChE in the group beta and the presence of AChE in the DC. There are some points of resemblance with the distribution of serotonin in the DAO and the caudal MAO of the cat, but a causal relationship between the presence of AChE and the distribution of certain afferent or efferent (peptidergic) systems of the olive has not been established. A dense plexus of ChAT-immunoreactive fibers pervading the entire inferior olive of the cat, was reported by Kimura et al. (1981). Receptor binding using labelled ~bungarotoxin for nicotinic receptors (Hunt and Schmidt, 1978) and [3H]propyl-benzilyl choline for muscarinic receptors (Rotter et al., 1979a) was stronger in the PO and the MAO, than in the DAO of the rat. Wamsley et al. (1981) and Swanson et al. (1987) reported on the presence of muscarinic and nicotinic receptors in the olive, but did not distinguish between the subdivisions of the inferior olive. 6.3.3. The connections between the inferior olive and the cerebellum

When the olivocerebellar projection was studied in the cat with antegrade axonal transport of tritiated aminoacids it appeared that the labelled climbing fibers were arranged 242

Ch.I

The cerebellum." chemoarchitecture and anatomy

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Fig. 169. Diagram of the distribution of AChE in the inferior olive of the cat. A. Transverse sections, numbered from caudally to rostrally. B. Diagrams of the unfolded inferior olive. Black: heavy staining; stippled: medium staining; white: low staining. Abbreviations: DAO = dorsal accessory olive; dc = dorsal cap; dmcc = dorsomedial cell column; MAO = medial accessory olive; nfl = subnucleus beta; PO = principal olive; vlo = ventrolateral outgrowth. Marani et al. (1977).

in longitudinal strips (Courville, 1975) and that this zonal pattern closely resembled the zonal organization in the corticonuclear projection and the associated compartmental subdivision of the feline cerebellum (Voogd, 1969; Groenewegen and Voogd, 1977; 243

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J. Voogd, D. Jaarsma and E. Marani

Groenewegen et al., 1979; Voogd and Bigar6, 1980). Electrophysiological studies of the olivocerebellar projection (Armstrong et al., 1973, 1974) and of spino-olivocerebellar climbing fiber paths (SOCPs) (Oscarsson, 1969, 1973, 1980) generally supported the zonal distribution of the climbing fibers. To express the similarity with the anatomical olivocerebellar climbing fiber zones Oscarsson and Sj61und (1977a) adopted a modification of Voogd's (1969) nomenclature to designate the zonal projections of the S O C P s . 4 Brodal and Kawamura (1980), who reviewed their extensive studies with retrograde labelling of the olivocerebellar pathway, similarly used Voogd's paradigm in the interpretation of their data. Information on the olivocerebellar projection in primates is less complete, but the available data suggest that its overall organization is similar to the cat. The principles underlying Voogd's nomenclature also have been applied to the olivocerebellar projection in the rat (Buisseret-Delmas, 1988a,b; Buisseret-Delmas and Angaut, 1993; Voogd, 1995) but in rodents other schemes have been proposed (Azizi and Woodward, 1987; Apps, 1990). The olivocerebellar projection to the anterior lobe, the hemisphere of the posterior lobe, the caudal vermis and the flocculus and the collateral projections to the cerebellar and vestibular nuclei will be reviewed. Some reports on the olivocerebellar projection in non-mammalian vertebrates are available. In the chicken the inferior olive consists of a dorsal and ventral laminae. The homologies of these laminae with the mammalian olive are complicated (see Furber, 1983; Arends and Voogd, 1989). The projection to the cerebellum is crossed and arranged in longitudinal zones (Freedman et al., 1977). A detailed topographical map of the relations between the olive and the cerebellum of the pigeon was published by Arends and Voogd (1989). Different lines of evidence on the myelo-architecture and the mossy and climbing fiber connections, therefore, support the longitudinal subdivision of the avian cerebellum. The inferior olive of the turtle is difficult to identify with histological staining methods. Ktinzle and Wiklund (1982) identified it on the basis of specific retrograde transport of D-[3H]aspartate. The olivocerebellar projection in the turtle also is organized in bands. Both in reptiles and birds the climbing fibers and the smooth parts of the Purkinje dendrites on which they terminate are limited to the deep parts of the molecular layer (compare the extent of the spiny branchlets in Purkinje cells of fish, birds and mammals in Fig. 12). 6.3.3.1. The olivocerebellar projection to the anterior lobe

The situation in the cat, where most data are available, will be considered first. The olivocerebellar projection to the anterior lobe in primates and the rat will be reviewed at the end of this section. Three broad zones, e.g. the vermis, the pars intermedia and the lateral zone or hemisphere proper, can be distinguished in the anterior lobe of the cat, in accordance with the classical description of Brodal (1940). The anterior vermis receives its climbing fibers from the caudal portions of the MAO and the DAO, the pars intermedia from the rostral MAO and DAO and the hemisphere from the PO. Voogd (1969), Armstrong et al. (1974) and Groenewegen and Voogd (1977) in the cat distinguished between a medial A zone, receiving fibers from the caudal half of the MAO and a lateral B zone which receives its climbing fibers from the caudal DAO (Fig. 119).

Zones defined by electrophysiological criteria usually are designated by lower case characters (i.e. the a, x, cl-3, d 1-2 zones; see Fig. 175).

4

244

The cerebellum." chemoarchitecture and anatomy

Ch. I

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Fig. 170. Laterality and somatotopical organization of the projection zones of the ventral funiculus spinoolivocerebellar climbing fiber path (VF-SOCP) in the cerebellum of the cat. A. Rostral aspect. B. Caudal aspect of the cerebellum. The position of the caudal border of the inferior colliculus (INF.COLL.) is indicated. Oscarsson and Sj61und (1977b).

Equivalent SOCP's terminating in the a and b zone of the anterior vermis, that could be activated from the isolated ventral and dorsal funiculus of the cord, were discovered by Oscarsson c.s. (see Oscarsson, 1969; 1973). The ventral funiculus (VF-) SOCP carries information from skin, joints and high and low threshold muscle afferents. The VFSOCP to the a zone is an ipsilateral pathway from the hindlimb. The VF-SOCP to the b zone is bilateral, the projection from the forelimbs was found to be located medially in the b (b~) zone, the hindlimbs in the more lateral b2 zone (Oscarsson and Sj61und, 1974, 1977a,b,c) (Figs 170 and 171). The a zone is also present as a wide, paramedian zone in lobule VIII; the b zone is restricted to the anterior vermis (Figs 170 and 183). The dorsal funiculus (DF-)SOCP synapses in the dorsal column nuclei and converges with the VF-SOCP at the level of the inferior olive. The DF-SOCP is activated from high threshold muscle afferents and from both tactile and nociceptive cutaneous afferents 245

Ch. I

J. Voogd, D. Jaarsma and E. Marani

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Fig. 171. Termination zones in the anterior lobe of the dorsal (A, DF-SOCPs), ventral (B, VF-SOCPs), dorsolateral (C, DLF-SOCPs) and lateral (D, LF-SOCP) spino-olivo-cerebellar paths. The diagrams represent those parts of lobules IV and V in the left half of the anterior lobe which are accessible at the cerebellar surface. Borders between zones a-d2 are indicated with lines. Areas activated from hindlimb and forelimb nerves through direct and indirect paths from the dorsal funiculus nuclei (DFN) or spinal cord to the inferior olive (IO) are indicated (see key). The DF-SOCP projection (from forelimb nerves) and the LF-SOCP projection to the c2 zone, and the VF-SOCP projection to the b zone are bilateral, whereas all other projections are ipsilateral. B-D based on Larson et al. 1969a, 1969b and Oscarsson and Sj61und 1977a, 1977b. Ekerot and Larson (1979a)

(Ekerot and Larson, 1979a; Ekerot et al., 1991a,b). Ipsilateral DF-SOCP's to the a zone and to the lateral b 2 zone, carry hindlimb information (Ekerot and Larson, 1979a; Ekerot and Larson, 1977, 1979a,b, 1982; see also Andersson and Erikson, 1981) de246

The cerebellum." chemoarchitecture and anatomy

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Ch. I

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CAMPBELL & ARMSTRONG'85 Fig. 172. Origin from the border region of the rostral and caudal medial accessory olive of the olivocerebellar projection to the electrophysiologically identified x zone in the cat. a = a zone; b = b zone; fl = subnucleus beta; cl-c3 = cl-c3 zones; coll = inferior colliculus; d~, d2 = dl and d 2 zones; dmcc = dorsomedial cell column; M A O = medial accessory olive; ml = mldline; pvg = lateral border of vermis; x = x zone; IV, V = lobules IV and V. R e d r a w n from Campbell and A r m s t r o n g (1985).

scribed a third, 'x' zone in the anterior lobe of the cat, located between the a and b zones (Fig. 171). The x zone is not present in all lobules of the anterior lobe, but is restricted to the lobules IV and V. It receives an ipsilateral projection through the DF-SOCP from the forelimb, through climbing fibers that branch between the x zone and zone in the pars intermedia (Fig. 175). The VF-SOCP does not terminate in the x zone. The climbing fibers to the x zone take their origin from the junction of the caudal and rostral halves of the MAO (Fig. 172) (Campbell and Armstrong, 1985; Voogd, 1989). Olivocerebellar fibers terminating in the A, X and B zones and the Purkinje cell fibers from these zones occupy the corresponding white matter compartments (Figs 119 and 120) (Voogd, 1969; Groenewegen and Voogd, 1977; Voogd and Bigar6, 1980; Voogd, 1989). The projections of the Purkinje cells of the VF-SOCP-innervated x and b zones were verified by Andersson and Oscarsson (1978a) and Trott and Armstrong (1987a,b) and were found to correspond to the anatomical corticonuclear projection zones X and B. The border between the B zone and the most medial C1 zone of the pars intermedia may be difficult to assess. This border was not recognized with retrograde labelling by Brodal and Walberg (1977a), who considered the B and C1 zone as a single area innervated by the caudal and lateral DAO. However, the presence of separate B and C1 compartments in the white matter and the characteristic electro-physiological properties of the b and c~ zones of Oscarsson validate their distinction in cat cerebellum. The problem in defining the border between the caudal portion of the DAO, projecting to the B zone and the more rostral regions, projecting to the C1 and C3 zones was discussed by Brodal and Kawamura (1980) and Voogd (1989). There is agreement on the differential origin of the two projections, but a precise border has never been established. Collateral projections to the fastigial and Deiters' nucleus take their origin from the 247

Ch. I

J. Voogd, D. Jaarsma and E. Marani

NOCICEPTIVECLIMBINGFIBER INPUT P

DAO DAO

A

1"I

MAO

~ULILr~TIVE NON-NOCICEPTIVE NOT STUDIED

Fig. 173. A summary diagram of nociceptive and non-nociceptive cutaneous climbing fiber input to lobules IV and V of the cerebellum of the cat. Forked arrows show branching of olivary axons to innervate pairs of zones (Ekerot and Larson, 1982). PF, primary fissure; DAO, dorsal accessory olive; MAO, medial accessory olive. Garwicz (1992).

olivocerebellar fibers to the A and B zones respectively (Groenewegen and Voogd, 1977). Retrograde labelling after injections of the fastigial nucleus of the cat was found in cells of the caudal MAO, the group beta and the dorsal cap (Hoddevik et al., 1976; Courville et al., 1977; Ruggiero et al., 1977). The projection of the lateral part of the caudal MAO to the lateral fastigial nucleus (Dietrichs and Walberg, 1985) may include the collateral projection of the olivocerebellar fibers to the x zone, that was never studied separately. Somatotopical patterns in the anterior vermis are arranged as mediolaterally disposed subzones or microzones. Information on the somatotopy of the A zone is scarce. According to Oscarsson and Sj61und (1977a,c) and Ekerot and Larson (1979a,b) the a zone is dominated by the hindlimb. The DF-SOCP projection to the x zone is dominated by the forelimb but lacks a somatotopical organization (Ekerot and Larson, 1977, 1979b, 1982). The exclusive forelimb connections of the x zone are in accordance with the cervical and cuneate projections to the border region of the caudal and rostral halves of the MAO (Boesten and Voogd, 1975; Gerrits et al. 1984a) that contains the olivary neurons projecting to the x zone (Campbell and Armstrong, 1985). Climbing fibers to the x zone receive tactile input and can be strongly activated by noxious pinch of the 248

The cerebellum." chemoarchitecture and anatomy I

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Fig. 174. Minimum threshold sites for evoking short latency climbing fiber responses from the pericruciate cortex of the cat in the b, c] and d, zones of the anterior lobe of cat cerebellum. ASG = anterior sigmoid gyrus; PSG = posterior sigmoid gyrus. Redrawn from Andersson and Nyquist (1983).

skin (Fig. 175). The receptive fields for tactile and noxious stimulation coincide (Garwicz et al., 1992 in Garwicz, 1992). Climbing fibers to the b zone are responsive to cutaneous and deep stimuli but do not receive a nociceptive input (Figs 170, 171 and 175). Successively more rostral body segments (tail, hindlimb, thorax, forelimb, head) project to successively more medial microzones in the b zone (Anderson and Oscarsson, 1978b; Andersson and Eriksson, 1981). The climbing fibers innervating a microzone share the same receptive field. In the b zone each microzone has been shown to project to a separate set of neurons of Deiters' nucleus (Andersson and Oscarsson, 1978a). The microzones and the olivocorticonuclear micro-complexes to which they belong can be considered as the basic functional units of the cerebellum. Contrary to the cortical zones to which they belong, the microzones do not possess anatomical borders because they are physiological artefacts isolated from a somatotopical continuum. Climbing fiber projections from the contralateral sensorimotor cortex to the lateral part of the anterior vermis were described by Provini et al. (1967, 1968). They noticed a strong correspondance in the localization of the cortical and peripheral projections to this region. Andersson and Eriksson (1981) and Andersson and Nyquist (1983) concluded that the projections of the posterior sigmoid gyrus to the a, x and b zones were closely matched to those of the VF-SOCP with respect to somatotopical organization and laterality. The projections to the a and b zones were bilateral and those to the x zone were crossed. Andersson and Nyquist (1983) identified short latency projections from separate forelimb and hindlimb areas in the posterior sigmoid gyrus to zones in the pars intermedia (Fig. 174). The short latency projections to the b zone took their origin from a single area, intermediate between and overlapping with the hindlimb and forelimb areas. Responses to stimulation of the more medial portion of this area, adjoining the localization of the hindlimb were located in the lateral b 2 zone, responses from more laterally situated sites, located next to the forelimb area, were located in the 249

Ch. I

J. Voogd, D. Jaarsma and E. Marani

medial b 1 z o n e . The x zone only received a short latency projection from the forelimb area. The subdivision of the pars intermedia into three zones, with the C1 and C3 zones receiving climbing fibers from the rostral two thirds of the dorsal accessory olive with collateral projections to the anterior interposed nucleus and the C2 zone from the rostral portion of the MAO with collateral projections to the posterior interposed nucleus (Fig. 119) (Groenewegen et al., 1979), agrees with the organization of the corticonuclear projection from these zones. C2 does not reach as far ventrally as the C1 and C3 zones and is absent from the lobules I and III, where the C~ and C3 zones become contiguous (compare Fig. 170). This arrangement agrees with other, anterograde and retrograde studies of the anterior lobe in the cat (Armstrong et al., 1974; Brodal and Walberg, 1977a; Kawamura and Hashikawa 1979; Gibson et al., 1987). It is generally assumed that the three anatomical C zones are identical to their electrophysiological namesakes identified as the terminations of the different SOCP's. The rostral segments of the c~ and c3 zones receive a converging short latency input from the ipsilateral hind-limb through the VF- and the DF-SOCP (Oscarsson and Sj61und, 1974; Ekerot and Larson, 1979a). Ipsilateral forelimb components of the DF-SOCP and of a pathway ascending in the dorsolateral funiculus (DLF-SOCP) (Larson and Oscarsson and Larson et al., 1969) converge upon the rostral segments of these zones. A third zone, receiving forelimb input through the DF-SOCP, was identified by Ekerot and Larson (1979a,b) in the extreme lateral part of the anterior lobe and indicated as the d 2 zone. The Cl, c3 and d2 zones receive branches from a common set of climbing fibers (Ekerot and Larson, 1977, 1982). One group innervates the medial halves of the Cl and the c3 zone, another set branches between lateral c3 and the d 3 zone, a third system of climbing fibers, gives off branches to the x zone and the lateral c~ (or cx) (Campbell and Armstrong, 1985) zone (Figs 171 and 173). The cl, c3 and d 2 z o n e s belong to a collection of zones, that also includes the x zone, receiving short latency input through the DF-SOCP. Moreover, these zones are distinguished from the intercallated b, c2 and d 1 zones, because they are strongly activated by nociceptive stimuli (Ekerot et al., 1991a,b; Garwicz et al., 1992, in Garwicz, 1992) (Fig. 175). The innervation of these zones from the inferior olive is not uniform. The medial Cl, the c3 and the d 2 z o n e are supplied by the rostral DAO, but the x and cx zones from the MAO (Fig. 172) (Campbell and Armstrong, 1985; Apps et al., 1991; Trott and Apps, 1991). As a consequence the anatomical

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Fig. 175. Correspondence between the classical three-zonal subdivision of the cerebellum of Brodal (1940: upper panel), the sagittal projection zones of the spino-olivocerebellar climbing fiber paths of Oscarsson c.s. (middle panel) and the anatomical zones of Voogd (lower panel). Arrows indicate the transverse branching of climbing fibers between zones (Ekerot and Larson, 1982). Hatched zones receive short-latency input from the DF-SOCP (Ekerot and Larson, 1979a) and are activated by nociceptive stimuli. Garwicz et al. (1992) in Garwicz (1992).

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Fig. 176. Somatotopical organization in the dorsal accessory olive (A,B) and the C3 zone (C) in the cat. A. Somatotopic organization in DAO. Each point summarizes the receptive fields of all cells recorded on a penetration at that point. Pooled data from 20 cats. Caudal to the right; medial side turned to bottom side diagram. B. Summarizes somatotopic organization. Dashed border between face and forelimb zones indicates that there were many forelimb cells in the face area. C. Topographical organization of the C , zone of the cat. Representation of nociceptive, climbing fiber projections of the forelimb. Cerebellar lobules are indicated to the left. Interrupted line indicates border between medial (right) and lateral (left) C, zone. Dig, digits; Lat, lateral; Vent, ventral; d, distal; p, proximal; r, radial; u, ulnar. A,B. Gellman et al. (1983); C. Ekerot et al. (1991a).

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J. Voogd, D. Jaarsma and E. Marani

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Fig. 177. Reconstructions of transverse sections through the anterior lobe from two experiments showing the

AChE banding pattern in the molecular layer, the responsive properties of the climbing fibers, and the locations of marking lesions. A. Transverse section through sublobules Vc to Vf showing a midline and bilateral lateral bands of AChE-positive activity. The area between the midline AChE band and the lateral border of the lateral band corresponds to zone A, as defined by Marani and Voogd (1977). All the units recorded within zone A were unresponsive to mechanical stimulation, except for one response that was located on the lateral edge of the lateral zone. Lateral to zone A, all the climbing fiber responses encountered represented areas of the extremities. B. Another experiment (similar to A except that the plane of the electrode penetrations and the histological section involved sublobules Vb to lobule III) where unresponsive units were encountered within zone A but lateral to the lateral AChE band; all of the climbing fiber responses were elicited by tactile stimulation. Robertson and Logan (1986).

and physiological nomenclatures for these zones no longer correspond. In an anatomical sense the cx (or lateral c~ zone), that receives climbing fibers from a caudomedial region of the rostral MAO, should be considered as part of the C2 zone. However, the physiological properties of the cx zone are indistinguishable from the (medial) c~ zone, which is innervated by the rostral DAO and represents the entire anatomical C1 (Fig. 173). A similar discrepancy may exist for the d 2 z o n e which, as the equivalent of the anatomical D 2 zone, supposedly receives climbing fibers from the dorsal leaf of the PO, but shares a climbing fiber projection with the c3 zone, that is innervated from the DAO. The intercallated c2 and d~ zones receive long latency projections through the DFSOCP, bilateral to the c2 zone, ipsilateral to d]. In addition a pathway in the lateral funiculus of the cord (LF-SOCP) (Larson et al., 1969b) provides a bilateral long-latency input to the c2 zone and the DLF-SOCP innervates the d] zone. Climbing fibers inner252

The cerebellum." chemoarchitecture and anatomy

Ch. I

vating the c3 and dl zones cannot be activated by nociceptive stimuli (Garwicz et al., 1992 in Garwicz, 1992) (Fig. 175). The multisynaptic pathways from the dorsal column to the inferior olive for these long-latency projections to c2 and dl, have not been identified. They may include connections from the dorsal column nuclei to the nuclei of the mesodiencephalic junction. The DF-SOCP projection to the c3 and presumably to the medial cl and d2 zones, is somatotopically organized. The rostro-caudal, hindlimb-forelimb organization of the pars intermedia has been known since Adrian (1943) and Snider and Stowell (1944). A corresponding rough topography in the projection of the lateral and rostromedial DAO to rostral and caudal segments of the C1 and C3 zones in the anterior lobe has been recognized with anatomical tracing methods (Fig. 119) (Brodal and Walberg, 1977a; Groenewegen et al., 1979; Gibson et al. 1987). The studies of Ekerot and Larson (1979b) and, especially, the mapping of nociceptive receptive fields of climbing fibers projecting to the anterior lobe by Ekerot et al. (1991a,b) showed a remarkable degree of somatotopical organization in the c3 zone (Fig. 176). The representation of the ipsilateral body half was double, with a mirror image of the sequence of receptive fields found in the medial and lateral c3 zones. A quite different, patchy and non-zonal type of somatotopical localization was described by Robertson (see Robertson, 1987 for a review), using natural stimulation in anaesthetized intact cats to record climbing fiber responses in the Purkinje cells. They found a relatively unresponsive area corresponding to the A zone, which was defined on the basis of AChE histochemistry (Fig. 177) (Robertson and Logan, 1986). A patch-like organization of the body representation exists throughout the anterior lobe hemisphere. Patches differ in size, isolated representations can be encountered within or adjacent to a patch involving a different area and adjacent patches seldom have representations of neighbouring skin areas. The somatotopical arrangement with forepaw in caudal and hindpaw in rostral parts of the anterior lobe is roughly maintained. The patchy representations in these areas resemble the 'fractured somatotopy' in mossy fiber systems (see Section 6.4.2.) and lack a definite sagittal zonal disposition. Branching of climbing fibers supplying the same sagittal zone, or a set of functionally similar zones, was first described by Armstrong et al. (1973). Systematic branching between the x and lateral c~, medial c~ and medial c3 and lateral c3 and d2 was described by Ekerot and Larson (1977, 1979a) and is illustrated in Figs 171,173 and 175. Branching between equivalent regions of the anterior lobe and the paramedian lobule has been anatomically substantiated by Brodal et al. (1980) and Rosina and Provini (1983) for the projections of the rostral DAO and MAO and for the PO of the cat, and by Eisenman and Goracci (1983), Payne et al. (1985); Wharton and Payne (1985) and Hrycyshyn et al. (1989) for the rat. Apps et al. (1991), who studied the branching between the x and cx zones of the cat cerebellum with double labelling methods, found only few doublelabelled neurons projecting to both zones in the border region of the rostral and caudal halves of the MAO. Branching to the medial c~, c3 and d2 zones was never studied with anatomical methods. Neurons supplying climbing fibers to the medial Cl zone are located in more lateral and caudal regions of the rostral DAO than those innervating the c3 zone (Trott and Apps, 1991). Neurons with branching axons should be located in the regions where the projections to c l and c3 overlap, and in the border region between the DAO and the ventral leaf of the PO, that innervates the D2 zone. It can be concluded that the anatomical and electrophysiological subdivisions of the pars intermedia of the anterior lobe of the cerebellum of the cat are not completely concordant (Fig. 173). Short-latency DF-SOCP innervated zones with cutaneous no253

Ch. I

J. Voogd, D. Jaarsma and E. Marani

ciceptive input alternate with zones lacking this input. This pattern extends into the vermis and the lateral zone. The short latency DF-SOCP zones are innervated from the MAO, rostral DAO and the PO. The lateral zone or hemisphere proper of the anterior lobe receives a projection from the principle olive. The distinction of this projection by Brodal (1940) was based on its absence in the cat, and its presence in the anterior lobe of the cerebellum of the rabbit, that extends further laterally. Projections of the PO to the extreme lateral part of the anterior lobe of the cat have been documented by Armstrong et al. (1974), Brodal and Walberg (1977a), Groenewegen et al. (1979) and Kawamura and Hashikawa (1979). Brodal and Kawamura (1980) discussed this projection and tentatively concluded that both the ventral and dorsal lamella of the PO projected to the anterior lobe, the dorsal lamella to the medial D1 zone and the ventral lamella to the lateral D 2 z o n e . It cannot be decided whether the dl and d 2 z o n e s identified in the electrophysiological studies of Ekerot and Larson (1979a, 1982) correspond to either D~ or the D2 zone. Some pertinent observations on the projection of the somatosensory cortex to the anterior lobe were made by Andersson and Nyquist (1983) (Fig. 174) in the cat. The c~, c2 and c3 zones receive short latency, somatotopically organized projections from the posterior sigmoid gyrus. The somatotopical organization of the c] and c3 zones is similar to that observed after peripheral stimulation, for the c2 zone no somatotopical organization was observed for the peripheral projections through the VF- and the DLF-SOCP. The corticocerebellar projection to the c2 zone is bilateral and the connections with the c~ and c3 zones are crossed, which is in accordance with the peripheral input to their zones. An exclusive projection to c2 was found to be present from the second somatosensory area. Short latency projections from the anterior sigmoid gyrus only terminated

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Fig. 178. Semidiagrammatic illustration showing the zonal arrangement in lobules IV and V of Saimiri based on projections from specific subnuclei of IO to each zone. A = A zone; B - B zone; Cl_3 - C].3 zones; Dr, 2 = DI,2 zones; caudmao = caudal medial accessory olive; daom3 = medial/lateral part of the dorsal accessory olive; dlpo = dorsal leaf of the principal olive; rostmao - rostral medial accessory olive; vlpo - ventral leaf of the principal olive. Whitworth and Haines (1986b).

254

The cerebellum. chemoarchitecture and anatomy

Ch. I

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Fig. 179. Diagram of lamellar and zonal distribution of olivary afferents and efferents in the rat. The two lamellae (folds) of the dorsal accessory olive (DAO, 1 and 2) and the horizontal lamella of the medial accessory olive (MAC, 3) appear to receive afferents mainly from the spinal cord and dorsal column nuclei while projecting to anterior vermis and parts of intermediate cerebellum. The medial MAC (vertical lamella, 4) receives afferents from the vestibular and visual areas and projects to the posterior vermis as well as the flocculus. The rostral lamella of MAC and both lamellae of the principal olive (PC) receive projections from higher centers and send fibers to the lateral hemispheres. In the lower part of the figure, three drawings of the inferior olive demonstrate the lamellae corresponding to their sagittal zones of projection in the cerebellum. Azizi and Woodward (1987).

bilaterally in the dl zone. This zone could also be activated from parietal cortical areas. The d 2 z o n e was not investigated, it seems likely that its cortical afferentation is similar to c~ and c3, i.e. a crossed, somato-topically organized input from the posterior sigmoid gyrus. 255

Ch. I

J. Voogd, D. Jaarsma and E. Marani

The organization of the olivocerebellar projection to the anterior vermis in primates appears to be similar to the cat (Brodal and Brodal, 1981; Whitworth and Haines, 1986b) (Fig. 178). Olivocerebellar projections to the A, X and B zones were identified in a preliminary report on Macaca fascicularis by Voogd et al. (1987a,b, 1990). The projection to the A zone resembles the situation in the cat with respect to the presence of A1 and A2 subzones. Olivocerebellar projections to the X zone take their origin from intermediate levels of the MAO, that also project to the C2 zone. A collateral projection to Deiters' nucleus detaches from olivocerebellar fibers to the B zone. Projections from the DAO and the dorsal and ventral leaf to the hemisphere of the anterior lobe were described by Brodal and Brodal (1981) in macaque monkeys and C1, C 2 and C 3 and D zones were identified by Whitworth and Haines (1986) in the olivocerebellar projection to the anterior lobe in Saimiri sciureus. Studies of the olivocerebellar projection in the rat included the anterior lobe (ChanPalay et al., 1977; Furber and Watson, 1983; Campbell and Armstrong, 1983; Azizi and Woodward, 1987; Buisseret-Delmas, 1988a,b; Buisseret-Delmas and Angaut, 1989b, 1993; Buisseret-Delmas et al., 1993). Olivocerebellar projection zones in the cerebellum of the mouse were studied by Beyerl et al. (1982). Of these studies the paper of Azizi and Woodward (1987) is of interest because it introduced a new scheme for the identification of the zones (Fig. 179). Azizi and Woodward (1987) subdivided the MAO in a horizontal lamella (i.e. the subnuclei a and b of the caudal MAO), a vertical lamella (including the subnucleus c, the group beta and the dorsal cap) and a rostral lamella (corresponding to the rostral MAO). They distinguished a dorsal fold of the caudal DAO, which is joined laterally to the rest of the DAO, that was indicated as the ventral fold. The different lamellae and folds were found to project in a systematic manner to sagittal zones (Fig. 179). In the anterior lobe they distinguished projections from the horizontal lamella of the MAO to a medial vermal zone (corresponding to the A zone) and from the dorsal fold of the DAO to a lateral vermal zone (corresponding to B). The ventral fold projects to a single zone in the pars intermedia (corresponding to C~) and the rostral lamella of the MAO to a zone in the lateral cerebellum (corresponding to C2). The dorsal lamella of the PO projects to a medial zone in the hemisphere (D~) and the ventral lamella of the PO to the most lateral zone of the hemisphere (D2). With respect to the situation in the cat, therefore, the projections of the dorsal and ventral leaf of the PO to the D~ and D2 zones of the anterior lobe of the rat cerebellum are reversed. The projection of the DAO to the C3 zone was not identified. Buisseret-Delmas (1988a and b), Buisseret-Delmas and Angaut (1989b, 1993) and Buisseret (1993) using the same retrograde transport methods as Azizi and Woodward (1987), identified projections to the A, X, B and C1-C3 zones from the same subnuclei of the inferior olive as in the cat. They confirmed the projection of the dorsal fold of the caudal DAO to the B zone and traced projections to the C1 and C3 zones from the rostrolateral and rostromedial DAO respectively (Fig. 141). The main differences between their scheme lobe and the situation in the cat are the the presence in the anterior lobe of the rat cerebellum of an additional Do zone and the reversal in the projection of the dorsal and the ventral leaf of the PO to the D~ and D2 zones. The olivocerebellar projection to the pars intermedia is similar in rat and cat. The differences in the connections of the C~_3zones of rat cerebellum concern the corticonuclear projection of the zones of the pars intermedia (see Section 6.1.4.). The X zone was indentified in the lobules IV-VI of the cerebellum of the rat by Buisseret-Delmas et al. (1993) as a zone located lateral to the A zone, projecting to the junctional region of the fastigial and interposed nuclei (their 'interstitial cell groups'). 256

The cerebellum." chemoarchitecture and anatomy

Ch. I

rostral

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1ram Fig. 180. Schematic illustration of the result of D-[3H]aspartate injection into lobules IV and V of the cerebellum of the rat. In the sketches of the cerebellar sections, retrogradely labelled axons and axon collaterals are indicated by lines and dots. Filled dots in the olives indicate the location of labelled cells. Retrograde labelling in cells of the inferior olive is also illustrated in more detail in the diagrams on the right. BP = brachium pontis; D A O = dorsal acessory olive; D N = Deiters' nucleus; F N - fastigial nucleus; LL = lateral lemniscus; LLV = ventral nucleus of the lateral lemniscusw; M A O - medial accessory olive; OI = inferior olive; OS = superior olive; PN - pontine nuclei; PO - principal olive; RB - restiform body; I-X lobules I-X. Wiklund et al. (1984).

It receives a projection from the subnuclei a, b and c of the caudal MAO. The origin of this projection in the rat appears to be more extensive than in the case of the X zone of the cat (compare Fig. 172) (Campbell and Armstrong, 1985). They also identified a CX zone in the lobules V and VI, medial to the C1, on the basis of its projection to the interstitial cell groups. It received its olivocerebellar projection from the group c at middle and rostral levels of the MAO. Buisseret-Delmas and Angaut (1989b, 1993) defined three D-zones in the lateral part of the anterior lobe on the basis of their corticonuclear projection to the dorsolateral hump (Do), the dorsal magnocellular part of the lateral cerebellar nucleus (D1) and the ventral parvicellular part of this nucleus (D2). They receive their olivocerebellar projections, respectively, from the DM group and the medial half of the ventral leaf of the PO (Do), the dorsal leaf of the PO (D,) and the lateral half of the ventral leaf of the PO (D2) (see Fig. 141). Several aspects of the olivocerebellar projection have been studied mainly in the rat. 257

Ch. I

J. Voogd, D. Jaarsma and E. Marani

D

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Fig. 181. A comparison of the zonal distribution of Zebrin-immunoreactive Purkinje cells (left diagram) and the olivocerebellar projection (right diagram) in the rat. Diagram of the olivo-cerebellar projection is based on data from Furber and Watson (1983), Eisenman (1984), Azizi and Woodward (1987), Buisseret-Delmas and Angaut (1989b, 1993), Sugita et al. (1989), Apps (1990), Akaike (1992) and Ruigrok and Voogd (1992). Subnuclei of the inferior olive (bottom diagram) and their projection zones are indicated with the same symbols. With the exception of the olivocerebellar projection to the lateral extension of the A zone (A~) of Buisseret-Delmas (1988a) (Figs 141 and 144), the correspondance between the Zebrin zones and the olivocerebellar projection has not been verified experimentally. 1-7 = Zebrin antigenic zones P1 - P7; a = subnucleus a of the MAO; A = zone A; A~ = lateral extension of the A zone of Buisseret-Delmas (1988a); b = subnucleus b of the MAO; B = zone B; 13 = group beta; c = subnucleus c of the MAO; C1-3 = zones C1-C3; CrI = crus I of the ansiform lobule; CrlI = crus II of the ansiform lobule; D = zone D; Do = zone Do of Buisseret-Delmas and Angaut (1989b); D A O = dorsal accessory olive; dc = dorsal cap; dl = dorsal leaf of the PO; D M = dorsomedial subnucleus of the ventral leaf of the PO; dmcc = dorsomedial cell column; F L O = flocculus; M A O = medial accessory olive; P F L = paraflocculus; P M D = paramedian lobule; PO = pricipal olivary nucleus; SI = simple lobule; vl = ventral leaf of the PO; vlo = ventrolateral outgrowth; X = zone X; I-X = lobules I-X.

The zonal distribution of the olivocerebellar projection was first demonstrated by ChanPalay et al. (1977), using autoradiography of antegrade axonal transport of 35S-methion258

The cerebellum." chemoarchitecture and anatomy

Ch. I

ine. She advocated a bilateral projection of the olive, but this observation has never been confirmed. The labelling of climbing fibers in the molecular layer in her experiments always was discontinuous. The unlabelled spaces between the labelled climbing fiber strips were assumed to receive climbing fibers of extra-olivary origin. This argument subsequently was disproved by Campbell and Armstrong (1983), who showed that labelling of climbing fibers from the inferior olive in the molecular layer was continuous. The investigations of Wiklund et al. (1984), using selective retrograde transport of [3H]D-aspartate were done in rats. The injection sites in the cerebellar cortex were relatively large and defied a detailed analysis of the material in terms of zones. Their experiments clearly confirmed the presence of branching olivocerebellar fibers between the anterior lobe and lobule VIII of the caudal vermis and the paramedian lobule. Moreover, their experiments clearly showed the presence of collateral projections to the cerebellar nuclei and to Deiters' nucleus (Fig. 180). The zonal disposition of Zebrin-immunoreactive and non-immunoreactive Purkinje cells has not been compared in any detail to the olivocerebellar projection. According to Gravel et al. (1987) some of the borders between Zebrin-positive and negative zones coincide with borders of certain climbing fiber strips, but these strips were not further identified. Judging from the reported identity of some of the Zebrin-positive and -negative zones with certain corticonuclear and cortico-vestibular projection zones it seems likely, that the correspondence between the zonal organization in the olivocerebellar projection and the Zebrin pattern will be close (Fig. 181). 6.3.3.2. Olivocerebellar projection to the hemisphere of the posterior lobe According to Groenewegen et al. (1979) the olivocerebellar projection zones in the cat continue uninterruptedly from the anterior lobe across the primary fissure into the simple lobule, where they diverge laterally to enter the ansiform lobule (Fig. 119). The projection of the caudal MAO to the B zone cannot be traced beyond the border of the simple lobule (lobule VI) and lobule VII. The projections from the rostral DAO to the C~ and C3 zones in the ansiform lobule are narrow or interrupted, but reappear in the caudal folia of the crus II (the ansula). C3 is only present in the dorsal folial rosette of the paramedian lobule, C1 continues into the most ventral folia of this lobule. The projection of the rostral MAO to the C2 zone can be traced as an uninterrupted zone, through the medial ansiform lobule, and the centrolateral paramedian lobule into the paraflocculus, where it occupies a ventral position in the dorsal paraflocculus and a dorsal position in the ventral paraflocculus. The final segment of the C: zone innervates the caudo-medial portion of the flocculus (Gerrits and Voogd, 1982). The PO is connected with the D~ and the D2 zones of the entire hemisphere and sends collaterals to the lateral cerebellar nucleus. The antegrade [3-H]leucine tracing experiments of Groenewegen et al. (1979) did not allow for an analysis of the possible differential origin of the climbing fibers terminating in the D1 and D2 zones. The olivocerebellar connections of the simple and ansiform lobules were studied in detail by Rosina and Provini (1982; see Kotchabhakdi et al. 1978, for an earlier report). Their conclusions are in accordance with the scheme of Groenewegen et al. (1979), specifying the presence of a C~ zone in the central axis of the folial rosette, and the attenuation of the C3 zone in the lateral bend of the lobule. They found a projection of the ventral leaf of the PO to the medial D1 zone and of the dorsal leaf to the lateral D 2 zone of the ansiform lobule (Fig. 182). The olivocerebellar projection to the crus II and the paramedian lobule of the cat was 259

Ch. I

J. Voogd, D. Jaarsma and E. Marani

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Fig. 182. The olivocerebellar projection to the lobulus simplex and the ansiform lobule of the cat. A. Diagrammatic surface view of the cerebellar injected areas in the four different groups of folia of cerebellum. Thin lines indicate H R P injected areas in the medial crus I-simple lobule; dotted lines in the lateral crus I; thick lines in the lateral crus II; dashed lines in the intermedio-medial crus II. Arrows indicate the subdivisions of the ansiform lobule adopted in this study: (a) intercrural sulcus and border between crus I and lateral crus II; (b) border between lateral and intermediate crus II; (c) intracrural sulcus 2 and border between intermediate and medial crus II. B. Summarizing diagram of the olivary areas retrogradely labelled by H R P injections into the four different groups of folia, drawn on the schema of the IO imagined unfolded. The olivary areas corresponding to the different groups of cerebellar injected folia are marked by thin, dotted, thick and dashed lines as indicated in A. Four different hatchings mark the four olivary subdivisions (MAO, vl, dl and lateral bend, DAO) which project to the four different longitudinal strips (C2, D1, D2 and C1-C3) of the ansiform lobule, as reconstructed in C. C. Surface view of the localization of the longitudinal zones as reconstructed by these reported results (D1 and D2 zones) and by previous studies (C2 and C1-C3 zones). Abbreviations: CrI, II = crus I, II of the ansiform lobule; C1-3 = C1-3 zones; D1,2 = D1,2 zones; D A O = dorsal accessory olive; dl = dorsal leaf of the PO; 1 = lateral; L.pm = paramedian lobule; L.sim = simple lobule; m = medial; M A O = medial accessory olive; Pfl.d = dorsal para-flocculus, PO = principal olive; vl = ventral leaf of PO. Rosina and Provini (1982).

the subject of series of reports, that reflect the changing views of the main actors in the course of time (Brodal and Courville, 1973; Courville et al., 1973; Brodal et al., 1975; Brodal and Walberg, 1977b; Walberg and Brodal, 1979). Brodal and Kawamura (1980), reviewing this work, concluded that C~, C2 and D zones are present in the paramedian lobule. They were unable to identify C3 in the dorsal lobules and stated that the dorsal and ventral leaf of the PO project to narrow D1 and D 2 z o n e s respectively. A reverse origin of the projection of the ventral and the dorsal leaf of the PO to the D~ and D 2 was advocated by Groenewegen et al. (1979) and substantiated by Rosina and Provini (1982) for the ansiform lobule. Electrophysiological studies of the configuration of the termination of climbing fibers in the paramedian lobule are scarce. Trott and Apps 260

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d

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C2 ci V

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| Fig. 183. Activation of climbing fibers projecting to the caudal paramedian lobule (PM) in (B) by stimulation of collaterals terminating in the c~, c2, c3 and d zones of the anterior lobe indicated in (A). Redwawn from Oscarsson and Sjolund (1977b).

(1993) identified c~, c 2 and c3 zones in the rostral paramedian lobule on the basis of their electrophysiologically defined climbing fiber input and their projections to the cerebellar nuclei. They concluded that the presence of the c3 zone was variable. Oscarsson and Sj61und (1977b) delimited an ipsilateral hindlimb VF-SOCP projection to cl and c3 zones in the pars copularis of the paramedian lobule: the zones fuse into a wide area occupying almost the entire ventralmost paramedian folium. Stimulation of the c1_3and the d zones in the anterior lobe activated climbing fibers in the corresponding zones of the paramedian lobule (Fig. 183). Jeneskog (1974) recorded climbing fiber evoked potentials in the c~ and d zones of the anterior lobe and the paramedian lobule, after stimulation in and around the red nucleus. Rubral and spino-olivary pathways converge upon these zones: the DLF-SOCP terminates in the d zone, and the hindlimb and forelimb components of the DF-SOCP in the ventral and dorsal portions of the Cl zone of the paramedian lobule (Jeneskog, 198 l a). Tentatively they identified a C3 zone in the dorsolateral paramedian lobule and located the C2 zone in the central region of this lobule after mesencephalic stimulation. The total picture of the olivocerebellar projection to the paramedian lobule is less complete, but still very similar to the anterior lobe, with fusion of the C~ and C3 zones in its ventralmost part. The discontinuity of the C3 zone in the dorsolateral and ventrolateral parts of the lobule allows the C2 zone to leave the paramedian lobule to enter the paraflocculus. 'The paraflocculus is probably the most enigmatic cerebellar lobule from a functional point of view' (Brodal and Kawamura, 1980), but the pattern of its climbing fiber 261

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J. Voogd, D. Jaarsma and E. Marani

afferents is simple and straightforward, with concentric projections of the rostral MAO to the C2 zone along its medial border and of the PO to the D zones along its periphery (Fig. 119) (see Brodal and Kawamura, 1980, for a discussion of the results of retrograde tracing studies from the paraflocculus). The main problem in the zonal structure of the paraflocculus is posed by the presence of a distinct subdivision of the D compartment and clear evidence for the projection of D~ and D 2 z o n e s to different parts of the dentate nucleus and the lack of evidence on a corresponding differentiation in the olivocerebellar projection from the PO. An educated quess can be made on the basis of the generally acknowledged nucleo-olivary projections of the rostromedial dentate to the dorsal leaf and the caudolateral dentate to the ventral leaf of the PO. Because the D~ zone projects caudolaterally and the D 2 z o n e of the paraflocculus rostromedially in the dentate, it seems likely that the D~ zone is innervated by the ventral leaf and the D 2 z o n e by the dorsal leaf of the PO. The olivocerebellar connections of the hemisphere of primates have been studied in the classical studies of Holmes and Stewart (1908), that were reviewed and updated by Jansen and Brodal (1958) (see also Voogd et al., 1990). Information on monkeys is limited to the retrograde transport experiments of Brodal and Brodal (1981, 1982) with injections of HRP in the simple, ansiform and paramedian lobules, in macaques, that suggest that C~, C2, C3 and D zones are present in these lobules. The diagram of Buisseret-Delmas and Angaut (1993) of the organization of the olivocerebellar projection in the rat (Fig. 141), appears very similar to the situation in the cat, but closer inspection learns that there are several important differences. One difference concerns the presence in the hemisphere of the posterior lobe of an additional zone, which has never been observed in the cerebellum of either cat or primates, located in the medial region of the hemisphere of the simple lobule, the ansiform lobule and the rostral folium of the paramedian lobule. This zone projects to the dorsolateral protuberance of the fastigial nucleus. It is absent from the anterior lobe and the copula pyramidis (see Section 6.1.4.). The olivocerebellar projection from the medial subnucleus c to this zone in the hemisphere (the lateral extension of the A zone of Buisseret-Delmas (1988a)) was recognized with retrograde transport from small WGA-HRP injections in this area (Fig. 142). She considered it to belong to and and to be continuous with the vermal A zone. The projection of the caudal MAO to the hemisphere was not noticed by Campbell and Armstrong (1983) and was not accounted for in Azizi and Woodward's (1987)

Fig. 184. The tecto-recipient zones in the vermis (lobule VII) and the hemisphere (paramedian lobule, Crus II and simple lobule) of rat cerebellum. Note their absence from anterior lobe, Crus I and copula pyramidis. Compare Fig. 142. ANT = anterior lobe; COP = copula pyramidis; CrI(II) = crus I(II) of ansiform lobule; PMD = paramedian lobule; SI = simple lobule. IV-IX = lobules IV-IX. Redrawn from Akaike (1992).

262

The cerebellum." chemoarchitecture and anatomy

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scheme of the olivocerebellar projection in the rat. Akaike (1986bc, 1987, 1989, 1992) identified a similar zone with stimulation of the ipsilateral superior colliculus. It receives climbing fibers from the rostral subnucleus c (the tectorecipient zone of the MAO), that branch between medial VIb and the crus II (Fig. 184). It differs from the lateral extension of the A zone of Buisseret-Delmas (1988a) because it is absent from the crus I and from the rostral folia of the paramedian lobule. According to Akaike (1986c) it is separated from a similar zone in the vermis of lobule VII that also receives a climbing fiber input from the tectum, by a narrow non-tectal zone which straddles the paramedian sulcus. The tectal zone of the hemisphere is bordered on its lateral side by a zone receiving climbing fiber input from the ipsilateral wishker area (Akaike, 1989). The olivary neurons projecting to the tectal zone of lobule VII are also located in the tectorecipient area of subnucleus c, medial to the neurons projecting to the tectal zone in the hemisphere (Akaike, 1986a). The two populations are completely separated and do not collateralize between vermis and hemisphere (Akaike, 1986a,b,c, 1992). The position of the lateral extension of zone A in the hemisphere of the simple lobule differs from Buisseret-Delmas' (1988b) diagram, because it is located lateral and not medial to the B zone (see Section 6.1.4.). This peculiar position was confirmed in an experiment with an injection of the antegrade tracer Phaseolus vulgaris lectin in the rostral subnucleus c of the rat (Fig. 144) (Voogd, unpublished), with climbing fiber labelling in this zone and a collateral projection to the dorsolateral protuberance of the fastigial nucleus. To make the picture complete, it should be remembered (Section 6.3.2.1.) that the rostral subnucleus c of the caudal MAO is the target of the nucleoolivary projection of the dorsolateral protuberance (Fig. 164) (Ruigrok and Voogd, 1990). Counterstaining with the Zebrin I antibody showed that the termination of the climbing fibers coincides with the area of the Zebrin immunoreactive zones P4b and 5a extending from the simple lobule into the paramedian lobule. C1, C2, C3 and D zones receiving climbing fibers from the rostral DAO and MAO and the PO presumably are present in more lateral parts of the hemisphere of the simple lobule. The crus I of the ansiform lobule seems to lack a tectal response zone (Akaike, 1986b, 1987, 1992), and B, C~ and C3 zones are not represented in the ansiform lobule (Furber and Watson, 1983; Buisseret-Delmas, 1988a,b). The lateral extension of the A zone (tectal response zone of Akaike) and the C1, C2 and C3 zones reappear in the crus II and the dorsal folia of the paramedian lobule (Furber and Watson, 1983; Buisseret-Delmas, 1988a,b). The lateral extension of the A zone (Buisseret-Delmas, 1988a) and the C3 zone are absent from the copula pyramidis, that contains representations of the C1, C2 and D zones (Fig. 185) (Eisenman, 1981; Azizi and Woodward, 1987; Apps, 1990). Multiple D zones with projections from the PO are present in the entire hemisphere. According to Azizi and Woodward (1987) and Buisseret-Delmas and Angaut (1989b) the medial D~ zone (zone 6 of Azizi and Woodward, 1987) and the lateral D2 zone (zone 7 of Azizi and Woodward), receive their climbing fibers from the dorsal and the lateral half of the ventral leaf of the PO respectively (Figs 141 and 179). The dorsolateral hump sends a nucleo-olivary projection to the dorsomedial subnucleus (DM) of the olive, that is attached to the ventral leaf of the PO and has often been confused with the DMCC (Azizi and Woodward, 1987; Ruigrok and Voogd, 1990). Olivocerebellar projections from this subnucleus to the hemisphere of the simple lobule, the crus II, the paramedian lobule and the copula, but not to the crus I, have been noticed by Furber and Watson (1983) and Buisseret-Delmas and Angaut (1989b, 1993: their Do zone) (Fig. 141). The projection of the DM to the Do zone, therefore, is limited to the same lobules as the projection of the DAO to the C~ and C3 zones. Little is known about the olivocerebellar 263

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J. Voogd, D. Jaarsma and E. Marani

B8 a

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caudal Fig. 185. The olivocerebellar projection to the pyramis and the uvula (lobules 8 and 9) of the rat cerebellum. A,B. Olivocerebellar projection zones of the lobules 8 and 9. C. Origin of these projections, indicated in diagrams of transverse sections through the inferior olive, a = subnucleus a of the medial accessory olive; b = subnucleus b of the medial accessory olive; beta = group beta; c = subnucleus c of the medial accessory olive; d - dorsal accessory olive; dm = dorsomedial subnucleus; rn - medial accessory olive; pr = principal olive; 8 and 0 = lobules VIII and IX of Larsell. Relabelled and reproduced from Eisenman (1984).

projection to the paraflocculus of the rat. According to Furber and Watson (1983) and Azizi and Woodward (1987) and Buisseret-Delmas and Angaut (1993) it receives climbing fibers from the rostral MAO and the PO. There is a good correspondence between the organization of the olivocerebellar projection in the rat and the cat. The main exceptions are the presence of the lateral extension of the A zone (or tectal response zone) and the Do zone in the hemisphere of 264

The cerebellum." chemoarchitecture and anatomy

Ch. I

the rat cerebellar hemisphere, that seem to be absent in the cat. The interruption of C1 and C3 zones in the crus I and the presence of a C3 zone in the crus II and the paramedian lobule, and its absence in the copula are in accordance with the findings in the cat. Projections of the dorsal and the ventral leaf of the PO to the D1 and D2 zone respectively, are still controversial in the cat, but seem to be well established for the rat. With the exception of the olivocerebellar projection to the lateral extension of the A zone, which coincided with the area of the P4b/P5a Zebrin immunoreactive Purkinje cell zones (Fig. 144), no direct comparisons of the olivocerebellar projection with the Zebrin pattern are available. However, in view of the similarity of the corticonuclear projection zones to the zonal arrangement in the olivocerebellar projection (see Section 6.1.4.), it seems likely that the P5b+ zone corresponds to C: and that the Zebrin-negative zones that border upon P5 correspond to the C~ and C3 zones (Fig. 181). 6.3.3.3. The olivocerebellar projection to the caudal vermis and the flocculus

In all species the zonal arrangement of the olivocerebellar projection to the vermis of the simple lobule (lobule VI) is very similar to the anterior lobe. Caudal MAO and DAO projections to A and B zones were traced to lobule VI in the cat (Groenewegen and Voogd) (1977) (Fig. 119) and the rat (Buisseret-Delmas and Angaut, 1993) (Fig. 141). The olivocerebellar projection to the rest of the caudal vermis differs substantially from that to the anterior lobe, with characteristic patterns for each of the lobules VII-X. Collateral projections to the fastigial nucleus have been observed with injections of antegrade tracers in the MAO and its subnuclei and labelling of (sub)zones in the caudal vermis (Groenewegen and Voogd, 1977), but these collaterals have not been systematically studied. Brodal and Kawamura (1980) and Sugita et al. (1989) noticed that the cells of the inferior olive projecting to the A zone of the lobules VI-VIII in cat and rat respectively, were arranged along the periphery of the caudal MAO, surrounding a central region projecting to the A zone of the anterior lobe (Fig. 186). The differences between their diagrams concern the origin of the projection to lobule VI, and the involvement of the group beta, which provides a sparse projection to lobule VII of the cat, (Hoddevik et al., 1976) but projects heavily to the lobules VI and VIII of the rat (Sugita et al., 1989; Apps, 1990). The A zone of lobule VI of the cat receives a single projection from a cell column in the medial caudal MAO, located lateral to the cells projecting to lobule VII (Fig. 186). The situation in the rat is more complicated with two labelled foci in the lobules Via and b after injections of WGA-HRP. One is located in the group beta, the other is a column situated along the lateral margin of the caudal MAO and extending along the border with the rostral half of this nucleus. The localization of the latter column is very similar to the cells projecting to the X zone of the anterior lobe in the cat (Campbell and Armstrong, 1985), and may correspond to the olivary neurons projecting to the X zone in the rat (Buisseret-Delmas et al., 1993). Retrograde labelling in the subnucleus c appears with injections of the caudal lobule VIC of the rat (Sugita et al., 1989). The existence of multiple sites in the caudal MAO that project to lobule VI of the rat is in accordance with the subdivision of this lobule in multiple corticonuclear projection zones and Zebrin-positive and -negative Purkinje cell strips (see Sections 5.6.1.3. and 6.1.4.). Laterally the A zone is bordered by the projection of the caudal DAO (cat: Hoddevik et al., 1976) or the dorsal fold of the DAO (rat: Buisseret-Delmas, 1988a; Azizi and Woodward, 1987) to the B zone. The B zone ends at the border of lobule VI and lobule VII in the cat, and at the VIa/VIb border in the rat. 265

Ch. I

J. Voogd, D. Jaarsma and E. Marani

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Fig. 186. The projection of the medial accessory olive to the vermis based upon retrograde tracing experiments in the cat. A ( 1 , 2 ) - A(1,2) zone; dm.c.col. = dorsomedial cell column; 1= lateral; m = medial; nucl. fl = subnucleus beta; I - X - lobules I-X. Brodal and Kawamura (1980).

According to Hoddevik et al. (1976) the main projection to lobule VII in the cat takes its origin from the rostromedial portion of the caudal MAO. This region receives afferents from the contralateral superior colliculus (Weber et al., 1978; Saint Cyr and Courville, 1982) and projects in a topical manner to the lobules VI and VII, with the rostral superior colliculus being located in the medial part of these lobules and the caudal colliculus more laterally (Kyuhou and Matsuzaki, 199 l a,b). An equivalent projection to lobule VII (the vermal visual area) was traced in macaque monkeys from the medial, Z-shaped portion of subnucleus b of the caudal MAO (Frankfurter et al., 1977; Brodal and Brodal, 1981; Yamada and Noda, 1987; Ikeda et al., 1989). The olivocerebellar projection to lobule VII of the rat differs from the adjoining lobules VI and VIII in being derived from a single cell column in the medial MAO (Sugita et al., 1989). This projection to lobule VII of the rat of the tectal response zone of the subnucleus c of the caudal MAO has been studied by Hess (1982b) and in a series of publications of Akaike (see Akaike, 1992 for a review). The neurons projecting to 266

The cerebellum." chemoarchitecture and anatomy

Ch. I

lobule VII are located in the caudal half of the tectorecipient zone of the caudal MAO, caudal to and intermingled with cells projecting to the lateral extension of the A zone in the medial hemisphere (see also the previous Section 6.3.3.2.). There is no climbing fiber branching between lobule VII and this zone in the hemisphere. The climbing fiber responses on stimulation of the superior colliculus occupy the medial two thirds of lobule VII (Akaike, 1985, 1986b,c, 1992). A similar, medial localization of the climbing fiber evoked potentials on tectal stimulation was reported in the earlier study of Jeneskog (1983) in the cat. The nature and the precise origin of the climbing fiber afferents to the lateral, non-tectal zone of lobule VII are not known. The area may be related to the X-zone or to a climbing fiber zone in more caudal lobules receiving afferents from the group beta. In the rat this non-tectal zone bridges the paramedian sulcus and consists of an 1 mm wide strip of cortex between the vermal and hemispheral zones supplied by the neurons of the tectorecipient area of subnucleus c of the caudal MAO. The relatively simple schemes of the olivocerebellar projection to lobule VIII based on studies in the cat by Hoddevik et al. (1976), Groenewegen and Voogd (1977) and Brodal and Kawamura (1980) now have evolved into the more complicated patterns defined by Eisenman (1981, 1984) and Apps (1990) for the rat. These patterns are of special interest because they bear a strong resemblance to the distribution of Zebrinpositive and -negative zones in lobule VIII of the rat (Hawkes and Leclerc, 1987) (see also Section 6.1.3.) and to the compartmentalization of this lobule in AChE-stained material of macaque monkeys (Hess and Voogd, 1986). Eisenman (1981) found two strips of climbing fibers in lobule VIII innervated by subnucleus a of the caudal MAO that were separated by a projection from the group beta. Climbing fibers terminating at the junction of lobule VIII with the copula pyramidis originated from the middle portion of the lateral DAO. Apps' (1990) observations were rather similar. The two strips receiving projections from subnucleus a constitute isolated regions, because projections from subnucleus a (and b: Sugita et al., 1989; Apps, 1990) are absent from lobule VII and scarce in IX. The group beta projection to lobule VIII, shifts medially in lobule IX, where it occupies a position next to the midline (Fig. 185) (Eisenman, 1984). This situation is reminiscent of the presence of distinct Zebrin-negative P 1 - and P 2 - strips, separated by P2+ in lobule VIII of the rat that disappear or become much narrower in the adjacent lobules VII and IX. The presence of an X zone in lobule VIII of the rat was postulated by Buisseret-Delmas et al. (1993). The olivocerebellar projection to lobule IX has been most extensively studied in cat and rabbit, data on rat and primates are less complete. Groenewegen and Voogd (1977) and Groenewegen et al. (1979) distinguished three zones in lobule IX of the cat: a medial zone, receiving a projection from the caudal MAO and/or the group beta, an intermediate zone receiving a projection from the DMCC and a lateral zone corresponding to C2, with climbing fibers from the rostral MAO. Similar patterns in the olivocerebellar projection to the uvula were reported by Brodal (1976) and Brodal and Kawamura (1980). Sato and Barmack (1985) in the rabbit and Kanda et al. (1989) in cat, distinguished projections of the group beta to two medial zones from the projection of the caudal MAO to a slightly more lateral area, and located a narrow strip receiving a projection from the ventral leaf of the PO between the lateral DMCC and rostral MAO (C2) zones (Fig. 187). Presumably, this pattern is shared by the rat (Eisenman, 1984; Bernard, 1987; Apps, 1990). Buisseret-Delmas et al. (1993) considered the projections from the DMCC and subnucleus c of the MAO to lobule IX of the rat to belong to the X and CX zones. Correlations of the olivocerebellar projection with the Zebrin pattern of lobule IX have not yet been verified. Retrograde labelling from lobule IX in the same 267

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subnuclei of the inferior olive was observed in the opossum (Linauts and Martin, 1978), in primates (Brodal and Brodal, 1981, 1982) and in subprimates (Whitworth et al., 1983). Zonal projections of the inferior olive to the nodulus (lobule X) and the flocculus have been substantiated in different species. Purkinje cells of these lobules are uniformly Zebrin-positive in rat, mouse, opossum, squirrel monkey (see Section 6.1.3.) and rabbit (Jaarsma unpublished observations). A compartmental subdivision of the white matter of the nodulus and the flocculus was demonstrated with AChE-histochemistry in the rabbit and the monkey (Section 6.1.5.) and correlated with the olivocerebellar projection to these lobules (Voogd et al., 1987a,b; Tan et al., 1995a,b). The pattern in the olivocerebellar projection to lobule X is quite distinct from lobule 268

The cerebellum." chemoarchitecture and anatomy

Ch. I

IX. The zones provided with climbing fibers from the group beta, the DMCC and the rostral MAO continue for some distance over the cortex of the nodule, but they are replaced by climbing fibers from the dorsal cap (DC) and the ventrolateral outgrowth (VLO). A large proportion of the climbing fibers from the latter two subnuclei are branches from climbing fibers which also terminate in the flocculus (Takeda et al., 1989a,b; Maekawa et al., 1989). Projections from rostral MAO, DMCC, group beta and DC, but not from the VLO were traced in the rabbit with HRP injections of lobule X and ventral IX by Alley et al. (1975). Zonally organized projections to lobule X from DC and VLO were demonstrated with anterograde axonal transport by Groenewegen and Voogd (1977) and Kawamura and Hashikawa (1979) in the cat. Retrograde labelling in these subnuclei after injections of retrograde tracers in lobule X was confirmed by Whitworth et al. (1983) for Galago and by Walberg et al. (1987) for the cat. Katayama and Nisimaru (1988) based their description of the olivocerebellar projection to lobule X on retrograde axonal transport experiments in the rabbit (Fig. 150). The projection from the group beta occupies the same medial position as in lobule IX. The lateral zones that are innervated from the DMCC and the rostral MAO, only extend over the dorsal surface of lobule X. The intermediate region contains a central zone, receiving a projection from the VLO, flanked by two strips innervated from the DC. The results of the anterograde and retrograde tracing experiments of Balaban and Henry (1988), also in the rabbit, were rather similar. They found the VLO zone to extend on the dorsal surface of lobule X and the ventral lobule IX and the two DC zones to be restricted to its ventral surface. These observations on the extent of the DC and VLO innervated zones are in complete accordance with the complex spike recordings in nodule Purkinje cells of the rabbit of Kano et al. (1990). The projections from the DC to lobule X and the flocculus were traced with parvalbumin-immunohistochemistry in rat pups by Wassef et al. (1992a,b). She showed that the cells of the DC (but not the cells of the VLO) transiently express immunoreactivity for antibodies against parvalbumin from birth till the 15th postnatal day. During this period two bundles of parvalbumin-immunoreactive climbing fibers were present in lobule X, corresponding to the medial and lateral DC innervated zones described in the rabbit. Olivocerebellar projections from the DC and the rostral one third of the MAO were first traced to the flocculus by Alley et al. (1975) with retrograde transport of HRP in the rabbit. The origin of the olivocerebellar projection to the flocculus was subsequently confirmed, using the same method, by Hoddevik and Brodal (1977), Yamamoto (1979) and Tan et al. (1995b) in the rabbit, Gerrits and Voogd (1982), Gould (1980) and Sato et al. (1983a) in the cat, Brodal and Brodal (1982) and Langer et al. (1985a) in macaque monkeys, Whitworth et al. (1983) in Galago and Blanks et al. (1983) and Bernard (1987) in the rat. The zonal organization of the olivocerebellar projection to the flocculus is similar in all species that were studied. Paired climbing fiber zones innervated by the caudal DC (the zones 2 and 4 of the rabbit flocculus) interleave with two zones innervated by the rostral DC and the VLO (zones 1 and 3) (Fig. 188). A fifth, lateral C2 zone is innervated by the rostral MAO. In the rabbit these zones correspond with the 5 corticonuclear and corticovestibular projection zones and the climbing fibers innervating a zone occupy the same white matter compartment as the axons of the Purkinje cells of this zone (Tan et al. 1995a,b,c) (Figs 147, 149 and 188) (see Section 6.1.5.). This arrangement is of special interest because it has been implicated in the spatial organization of eye movement control. Briefly the caudal DC of the rabbit transmits information about contralateral 269

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Fig. 188. Diagram of the origin of the climbing fiber afferents and the projections of the Purkinje cells of the five Purkinje cell zones of the flocculus of the rabbit. The zones and the corresponding white matter compartments containing their Purkinje cell axons and climbing fiber afferents are indicated with the same symbols. 1-4 = compartments and zones 1-4; DC = dorsal cap; fl-4 = folia 1-4 of the rabbit flocculus; fm = medial folium of the flocculus; fp = folium p of the ventral paraflocculus; Ip = posterior interposed nucleus; M A O = medial accessory olive; MV = medial vestibular nucleus; SV = superior vestibular nucleus; VLO = ventrolateral outgrowth; Y = group y. From Tan et al. (1995b,c).

movements of the surround around a vertical axis. Cells in the rostral DC and the VLO are excited by movements around an oblique horizontal axis that is oriented roughly parallel to the axis of the ipsilateral anterior semicircular canal (see Van Der Steen et al., 1994 for a review). Groenewegen and Voogd (1977) first described the projection of the caudal DC to a central zone in the cat flocculus flanked by two zones innervated by the rostral DC/VLO (zones 2 and 4). This pattern was extended by Gerrits and Voogd (1982) in tracing studies in the cat. The 7 floccular zones distinguished by these authors correspond to the 5 zones of the rabbit flocculus, because two of them are double (Fig. 149). Moreover, Gerrits and Voogd (1982) noticed that the zonal pattern in the olivo270

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cerebellar projection extends beyond the posterolateral fissure in the ventral paraflocculus (the 'ME': the medial extension of the ventral paraflocculus) (Fig. 149). In the anterograde axonal tracing studies of Tan et al. (1995b) in the rabbit the olivocerebellar projection was matched with the white matter compartmentalization in adjacent AChE-stained sections and, consequently, with the corticovestibular and corticonuclear projection of the correspooding zones (Fig. 188) (see Section 6.1.5.). The medial and central compartments 4 and 2 contained fibers from the caudal DC and the more laterally located compartments 3 and 1 conducted fibers from the rostral DC and the VLO to corresponding zones in the molecular layer. The fifth, most lateral compartment contained fibers of the rostral MAO to the Cz zone. The rostral DC and VLO 271

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Fig. 190. Distribution of C R F and CGRP-immunoreactive climbing fibers in P7 mouse cerebellum (A-C) and in neurons of the inferior olive (D-F). CRF-immunoreactive climbing fibers and neurons are indicated by dots, CGRP-immunoreactive climbing fibers and neurons by open circles, a, b and c = subnuclei a, b and c of the medial accessory olive; beta + group beta C1, C2 = Crus I and II; D A O - dorsal accessory olive; dc = dorsal cap; E G L . M L = external granular layer and molecular layer; F -- flocculus; I G L = internal granular layer; PF = paraflocculus; M A O = medial accessory olive; PO = principal olive; PL - Purkinje cell layer. Redrawn from Yamano and Tohyama (1993). (

projections extended, across the posterolateral fissure in folium P of the ventral paraflocculus, where the compartments 1 and 3 fuse around the tip of compartment 2, containing the olivocerebellar fibers of the caudal DC. The olivocerebellar projection to the flocculus of primates was not yet studied in detail. Zone 2 of the flocculus of macaques receives a projection from the caudal dorsal cap and, therefore, corresponds to zone 2 of the rabbit flocculus (Fig. 149). Recently Ruigrok et al. (1992) analysed the olivocerebellar projection to the flocculus and the adjacent paraflocculus in the rat with anterograde axonal tracing with the lectin Phaseolus vulgaris and retrograde transport of WGA-HRP (Fig. 189). Their results confirmed and extended the observations in other species, discussed in the previous paragraphs (Fig. 149). Two pairs of interdigitating zones, innervated by the caudal DC and the rostral DC/VLO, and a lateral C2 zone could be distinguished in the rat. The distal segments of these zones crossed the posterolateral fissure and were found to receive climbing fibers from more rostromedial levels of the DC (the FE/FE' zones, corresponding to zones 2 and 4 of the rabbit) and the ventral leaf of the PO (the FD'/FD zones, corresponding to zones 1 and 3). The rostral shift in the origin of climbing fibers directed to more distal (rostral) segments of a set of continuous, olivocerebellar projection zones is in accordance with Yamamoto's (1979) observations on the projection of intermediate levels of the DC and of the PO to the folium P of the ventral paraflocculus of the rabbit, and with Gerrits and Voogd's (1982) distinction of a lateral and rostral shift in the projection of successively more rostral parts of the DC and the VLO/PO to flocculus and ventral paraflocculus of the cat. Collateral projections of the olivocerebellar pathways from the DC and the VLO to the medial and superior vestibular nuclei were described by Balaban (1984), but have been denied by Groenewegen and Voogd (1977), Gerrits and Voogd (1982) and Ruigrok et al. (1991). Ruigrok et al. (1992) observed a strong projection to the parvocellular lateral cerebellar nucleus concomitantly with the climbing fiber labelling in the FD'/D zones. The anatomical and electrophysiological studies of Takeda and Maekawa (1989a and b), Maekawa et al. (1989), Kusonoki et al. (1990) and Kano et al. (1990, 1991) in the rabbit have shown that 36% of the climbing fibers innervating the flocculus and 64% of the climbing fibers in the nodule that could be driven by optokinetic stimuli, could be activated by collaterals terminating in the complementary lobule of the vestibulocerebellum. Retrograde labelling from flocculus and nodule with different fluorescent tracers resulted in double labelling of 9-27% of the DC neurons and 12-48% of the cells of the VLO. There was no distinct spatial segregation of the cells projecting to the two lobules.

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Fig. 191. Schematic line drawings of the unfolded opossum cerebellum modified after Larsell and Jansen (1972). The broken lines indicate the boundaries of the corticonuclear zones A-D after Klinkhachorn et al. (1984a). The distribution of the three types of enkephalinergic axons is indicated by the frequency and size of the symbols: the beaded axons by asterisks (C), the mossy fibers by dots (A), and the climbing fibers by triangles (B). I-X indicate vermal lobules; CR I, II, crura I and II, F, flocculus; LS, lobulus simplex; PFL, paraflocculus; PML, paramedian lobule. D. Distribution of enkephalinergic axons in a horizontal section through the cerebellar nuclei. D, dentate nucleus, F, fastigial nucleus; IPA, anterior interposed nucleus; IPP, posterior interposed nucleus. From King et al. (1987). (

6.3.4. The distribution of peptides and calcium binding proteins in climbing fibers and cells of the inferior olive A transient immunoreactivity in subpopulations of climbing fibers during the maturation of the cerebellum was described for somatostatin and CGRP. Somatostatin-immunoreactive climbing fibers are located in midsagittal and parasagittal bands in the vermis of lobules VI-VIII, some of which remain present in the flocculus and paraflocculus of adult rats (Vincent et al., 1985; Villar et al., 1989) (see Section 3.1.3., Fig. 20). Neuronal labelling in the inferior olive was limited to the MAO. Transient immunoreactivity for CGRP in immature climbing fibers, surrounding the somata of Purkinje cells, and in cells of the inferior olive was noticed in the early postnatal period in rats and mice (Kubota et al., 1987, 1988; Morara et al., 1989, 1992; Rosina et al., 1992; Chedotal and Sotelo, 1992; Yamano and Tohyama, 1993, 1994). These immunoreactive climbing fibers define longitudinal bands, first next to the midline, later in paravermal regions, the hemisphere and in the flocculus. Chedotal and Sotelo (1992) used the expression of CGRP immunoreactivity during prenatal development in the rat to trace olivocerebellar fibers to the cerebellar anlage in the fetal rat at El7. They identified the transient CGRP-immunoreactive climbing fibers in early postnatal stages bands as the A, B and C3 zones. Yamano and Tohyama (1994) noticed in early postnatal mice, that the bands of CGRP-immunoreactive climbing fibers alternate with CRF-immunoreactive climbing fibers (Fig. 190A-C). In postnatal mice CGRP disappears from the climbing fibers and the number of climbing fibers expressing CRF increases. Immunoreactivity for CGRP and CRF was never observed in the same neurons of the inferior olive. Both CGRP and CRF containing neurons occur in the group fl, subnucleus c of the MAO, the dorsal fold of the DAO, and in the dorsal cap (Fig. 190D-F), but the two cell populations occupy slightly different regions of these subnuclei. Their results on the localization of alternating bands of CRF and CGRP-immunoreactive climbing fibers in lobule X resemble the identification of parvalbumin-immunoreactive climbing fibers in two dorsal cap-innervated stripes in this lobule of the immature rat cerebellum by Wassef et al. (1992a,b) (see also Section 6.3.2.1.). The results of Yamano and Tohyama (1994), however, are difficult to interpret in terms of olivocerebellar projection, because their definition of the olivary subnuclei in the mouse is not precise enough. Enkephalin (ENK)-like immunoreactivity in subpopulations of climbing fibers and cells of the inferior olive has only been found in the opossum (King et al., 1986a, 1987) but not in other adult species. A more wide-spread distribution of corticotrophin releasing factor (CRF)-like immunoreactivity in certain climbing fibers has been observed in the cerebellum of the opossum (Cummings et al., 1989) and several other species including rat and mouse (Cummings et al., 1983; Van den Dungen et al., 1987), rabbit (Errico 275

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and Barmack, 1993), cat and sheep (Kitahama et al., 1988; Cummings et al., 1988, 1989), and Saimiri sciurus and Macaca fascicularis (Foote and Cha, 1988; Cha and Foote, 1988). The distribution of CRF, ENK and CCK in the cerebellum and the precerebellar nuclei, the species-differences and the co-localization of these peptides, recently was reviewed by King et al. (1992). Climbing fibers containing ENK are located in patches, that constitute discontinuous 276

The cerebellum." chemoarchitecture and anatomy

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midline and parasagittal bands in the vermis of opossum cerebellum (Figs 191B and 192) (King et al., 1986a, 1987). The position of the parasagittal band in the anterior lobe approximately corresponds to the border of the A and B zones, as determined by Klinkhachorn et al. (1984a and b) from the corticonuclear projection in this species. This description also would fit a localization in the X zone, which has not yet been described in the opossum. The presence of immunoreactive somata in the lateral cell group a of the caudal MAO (Walker et al., 1988; King et al., 1989) is in accordance with such an enkephalinergic climbing fiber projection to the X-zone. ENK-immunoreactive climbing or mossy fibers have not been observed in the cerebellum of rat, cat or primates. However, opiate receptor binding is present over the molecular and to a lesser extent over the granular layer of rat cerebellum (Zajac and Meunier, 1980; Robson et al., 1984). A detailed description of the localization of CRF in climbing fibers is available for the opossum (Cummings et al., 1989). CRF is present in all lobules but the numbers of reactive climbing fibers and of CRF-like immunoreactivity in individual fibers show local differences. CRF-immunoreactive climbing fibers are concentrated in midline and parasagittal bands in the vermis (Fig. 193), the density of the CRF climbing fiber innervation is high for the vermis of the lobules VI-VII and lobule X and the flocculus. Concentrations of less reactive climbing fibers occur in the hemisphere. CRF-containing climbing fibers are concentrated in the flocculus and the adjacent part of the paraflocculus. The density of CRF-immunoreactive climbing fibers and the density of CRF receptors, determined by radioligand binding of [125I]-Tyr-ovine CRF, show a close correspondence for this region (Cummings et al., 1989). The distribution of CRF in neurons of the inferior olive of the opossum was determined with immunohistochemistry and in-situ hybridization (Cummings et al., 1989). All subdivisions of the olive contain CRF-positive neurons, but the number of labelled neurons and their staining intensity varies within and among these subnuclei. Staining levels were consistently lower in subnucleus a of the caudal MAO, when compared to the subnuclei b and c (which includes the neurons of the less immunoreactive group beta). Cells in the dorsal cap and the rostral MAO stained intensely with both methods. Rostral DAO, especially its lateral part, contains more labelled cells than medial and caudal regions of this subnucleus. Cell bodies in the PO reveal less CRF immunoreactivity, but the caudal PO contains strongly immunoreactive neurons. In the opossum CRF-like and ENK-like immunoreactivity co-exist in climbing fibers in midsagittal and parasagittal bands at the border of the lobules VII and VIII, from here they shift to a position in the base of the paramedian lobule, laterally in lobule X and ventrally in the flocculus. CRF is more widely distributed in the climbing fibers and single-labelled and double-labelled climbing fibers co-exist in these foci. CRF-like immunoreactive neurons are double-labelled for ENK in the subnuclei a and c of the caudal MAO and in the dorsal cap (Cummings and King, 1990). This pattern corresponds with the known projections of the periphery of the caudal MAO (subnuclei a, c and group beta) to the caudal vermis and of the dorsal cap to a lateral zone in lobule X and to the flocculus and the adjacent paraflocculus (see Section 6.3.3.3.). A few CCK-labelled climbing fibers are located in the same region of the caudal vermis of the opossum as the ENK/CRF (double) labelled fibers. They may originate from CCKpositive neurons in subnucleus c, where most cells are also irnmunoreactive for CRF (King and Bishop, 1990). The widespread distribution of CRF-containing climbing fibers in the cat resembles the situation in the opossum (Cummings et al., 1988; Cummings, 1989). Some of the CRF in climbing fibers is concentrated in bands. Bands are prominent in midline and 277

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Fig. 193. Camera lucida drawings of 60/lm transverse sections (A,B) through the opossum cerebellum that illustrate the distribution of corticotrophin releasing factor immunoreactive (CRF-IR) climbing and mossy fibers. Stripes in the molecular layer represent climbing fibers; the dots represent mossy fibers. The density and intensity of each symbol represent the relative number of fibers and staining intensity for respective fibers within each region. Roman numerals I-X indicate cerebellar lobules. PML, paramedian lobule~ LS, lobus simplex; CRII, crus II; PF[, para-flocculus; F, flocculus; DN, deep nuclei; x7. C. Camera lucida drawing of a 60 pm sagittal section of vermal lobule X. M, molecular layer; P, Purkinje cell layer; G, granule cell layer; pc, Purkinje cell; mf, mossy fibers; CF, climbing fibers. Note greater density of mossy fibers at the ventral medial aspects of folium. Bar in A,B = 1 ram, in C = 70 pm. Cummings et al. (1989). <.-

paramedian regions of the anterior lobe, in the intermediate part of the anterior lobe and the simple lobule, in the caudal Crus II, the paramedian lobule and the caudal vermis (Fig. 194). The distribution of CRF-containing climbing fibers in the flocculus and the adjacent ventral paraflocculus is heterogeneous, CRF-like immunoreactivity is less prominent in the rest of the paraflocculus. When we compared the immunoreactivity to the distribution of AChE in the anterior lobe, the paramedian band of CRF-containing climbing fibers appears to coincide with the lateral border of the paramedian AChE-positive band in the molecular layer, i.e. with the X-zone, and/or the lateral A-zone. The same position was found for the paramedian band of CRF-containing climbing fibers in the opossum. The two prominent bands of CRF-containing climbing fibers in the intermediate zone probably are located within the C1 and the C2 zones, the C3 zone seems to be less immunoreactive. CRF-containing cell bodies in the inferior olive of the cat occur in all subdivisions, with a columnar distribution in medial and lateral parts of the caudal MAO, CRF-positive cells in the groups beta, the dorsal cap and the ventromedial cell column, and a predominant localization in caudal and lateral D A O (Kitahama et al., 1988; Cummings, 1989). All cells of the inferior olive of the monkey express some degree of CRF-immunoreactivity (Cha and Foote, 1988), activity is highest in the caudal and central MAO. CRFcontaining climbing fibers in the cerebellar cortex are present in most of the cortex. A high and uniform distribution is present in lobule IX, dorsal VIII and in the ventral paramedian lobule (the authors identified this lobule as belonging to the paraflocculus) and as midline paramedian and mid-hemispheral bands in the A and C2 zones of the anterior lobe (Figs 195 and 196). The localization in the C2 zone would be in accordance with the situation in the cat. The zonal distribution in the ventral anterior lobe is more complicated. CRF-receptor binding, using [~2sI]-ovine C R F was higher over the molecular layer (Millan et al., 1986). In the human brain immunoreactivity was present in the great majority of the neurons of all subdivisions of the inferior olive and in climbing fibers in the molecular layer of the anterior vermis (Powers el al., 1987). Originally immunoreactivity with an antibody against C R F could not be detected in rat cerebellum (Swanson et al., 1982). C R F immunoreactive fibers in the molecular layer were observed by Olschowka et al. (1982), Cummings et al. (1983), Merchenthaler (1984), Sakanaka et al. (1987) and Van den Dungen et al. (1987, 1988). Palkovits et al. (1987) identified these fibers as climbing fibers with light and EM-immunohistochemical methods. They found CRF-immunoreactive neurons in all subdivisions of the inferior olive with a predominance in the PO. C R F m R N A was transcribed by the cells of the inferior olive and depleted at'ter a lesion of the contralateral olivocerebellar tract. C R F receptor binding, using [I~25]-ovine CRF, was highest over the granular layer of rat cerebellum (DeSouza et al., 1985). It is not possible to correlate the distribution of immunoreactive cells in the adult 279

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inferior olive with the olivocerebellar projection more precisely without double labelling experiments of olivocerebellar fibers with CRF-immunocytochemistry. It seems likely, however, that CRF-immunoreactivity is not confined to certain subsystems and that gradients in the different subdivisions of the inferior olive are responsible for the banded 281

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J. Voogd, D. Jaarsma and E. Marani

Fig. 196. Dark-field photomicrograph of a sagittal section through the vermis of the cerebellum of the squirrel monkey. Note that some areas of the molecular layer (M) contain dense CRF-immuno-reactive axons while others do not. However, intensely immuno-reactive axons are evident in the Purkinje cell layer (P) even in those regions in which such axons are not evident in the molecular layer. Occasional labelled axons are evident in the granular layer (G) and in the white matter (W). Scale bar = 100/lm. Cha and Foote (1988).

distribution in the cortex. Interestingly, the a m o u n t of C R F - l i k e i m m u n o r e a c t i v i t y in cells of the inferior olive increases in pontine cats ( K i t a h a m a et al., 1988). C R F messenger R N A m e a s u r e d with a [35S]-labelled oligonucleotide probe increased in neurons of the caudal dorsal cap of rabbits after long-lasting optokinetic stimulation of the contralateral eye ( B a r m a c k and Young, 1990) (Fig. 197). By itself C R F has little or no effect on n e u r o n a l activity, but it potentiates the excitatory effects of g l u t a m a t e and aspartate, the putative n e u r o t r a n s m i t t e r s of the climbing fibers on the Purkinje cells (Bishop, 1990; Bishop and K e r r 1992). K i t a h a m a et al. (1988) also raised the question whether the C R F in precerebellar nuclei is under endocrinological control, but they were unable to observe effects of a d r e n a l e c t o m y or hypophysectomy. Some calcium-binding proteins occur in subpopulations of climbing fibers. Transient labelling with antibodies against p a r v a l b u m i n was observed in neurons of the dorsal cap

Fig. 197. Optokinetically induced increase in corticotrophin releasing factor (CRF) mRNA in caudal dorsal cap of the inferior olive of the rabbit revealed by darkfield photomicrograph of an emulsion-coated brain-stem section. The rabbit received 37 hr of binocular optokinetic stimulation in the posterior to anterior direction with respect to the left eye, causing a 360% increase in levels of CRF mRNA in the right dorsal cap. A,B. Bright-field and dark-field views of the same tissue section are shown. The finer spatial resolution of the emulsion demonstrates clustering of silver grains over individual olivary neurons. Scale bar = 200 r (Barmack and Young, 1990) 282

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and in climbing fibers in the two DC-innervated zones in lobule X and in a single zone in the rat flocculus by Wassef et al. (1992a,b) in rat pups. Calretinin-immunoreactive climbing fibers were observed in the cerebellum of the chicken (Rogers, 1987) and the rat (Floris et al., 1994). The precise topography of these climbing fibers has not yet been reported. 9 ~.-.

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6.4. MOSSY FIBER SYSTEMS

6.4.1. Concentric and discontinuous, lobular arrangements of mossy fiber systems Mossy fibers take their origin from many different sites in the brain stem and the spinal cord. The topographical organization of mossy fiber systems differs considerably from the climbing fiber projection. Mossy fiber systems enter the cerebellum through the restiform body and the middle cerebellar peduncle, lateral to and along the brachium conjunctivum. The main orientation of the stem fibers of the mossy fibers is transverse. These stem fibers are known as semicircular fibers, which constitute a layer situated rostral and dorsal to the cerebellar nuclei and peripheral to the olivocerebellar fibers, which are immediately apposed to the surface of the cerebellar nuclei and their efferent tracts (Fig. 198). Medially the layer of semicircular fibers continues into the cerebellar commissure, that consists of a central part, containing the decussation of the uncinate tract and the more peripherally located decussation of the mossy fibers (Voogd, 1964). Most mossy fiber systems are partially crossed. Secondary vestibulocerebellar and ventral spinocerebellar fibers, that enter the cerebellum in a ventral and medial position, occupy a ventral and rostral position in the cerebellar commissure; pontocerebellar fibers that enter the cerebellum laterally and caudally, occupy a dorsal and caudal position in the commissure. These fibers are located in the dorsal white matter of the anterior lobe, in the medullary ray of the lobules VI and VII and in the base of lobule IX. On their way towards the commissure the pontocerebellar fibers come to the meningeal surface, in the bottom of the intercrural fissure, where cortex is absent in the center of the ansiform lobule and in the interparafloccular sulcus in the center of the parafloccular loop, where the cortex is also interrupted. Mossy fiber systems within the cerebellum display a concentric arrangement: vestibulocerebellar fibers are located most centrally, and terminate in the cortex in the bottom of the fissures; spino-, cuneo-, and reticulocerebellar fibers extend more peripherally and

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pontocerebellar fibers cover the apex of the lobules. An electrophysiological corrolary of this concentric arrangement is the observation of superficial and deep terminations of exteroceptive and proprioceptive components of the cuneocerebellar tract respectively (Ekerot and Larson, 1972). As a consequence mossy fiber projections are discontinuous in the direction across the folium: being absent at the base or apex of the folium, and reappearing in the next (Jasmin and Courville, 1987b; Tolbert et al., 1993). Another consequence is that the distribution of mossy fibers is mainly transverse and lobular, with the restriction that these systems generally populate several adjacent lobules and that the borders between different mossy fiber systems do not necessarily correspond to the bottom of the fissures. The presence of a concentric arrangement in their distribution does not preclude a large degree of overlap between different mossy fiber systems. The question of the presence of overlap in their terminations is complicated by the termination of some mossy fiber systems in alternating longitudinal strips. The origin and distribution of primary and secondary vestibulocerebellar mossy fibers has been studied repeatedly. Vestibular root fibers enter the cerebellum from the superior vestibular nucleus, as part of the ascending branch of the vestibular root (Mannen et al., 1982; Sato et al., 1989). The development of the primary vestibulocerebellar projections was studied in rat embryos, where the root fibers can be distinguished by their parvalbumin-immunoreactivity (Morris et al., 1988). The literature on the projection of the vestibular nerve was reviewed in the study of Gerrits et al. (1989) on the primary vestibulocerebellar projection in the rabbit. The evidence on the origin and distribution of secondary vestibulocerebellar projections was reviewed for the cat by Brodal (1974), Kotchaphakdi and Walberg (1978), Batini et al. (1978, lobules VI and VII), Matsushita and Okado (1981, lobules I and II), Sato et al. (1983b, flocculus), Magras and Voogd (1985) and Blanks (1990, flocculus), for the monkey by Brodal and Brodal (1985), Langer et al. (1985a, ftocculus) and Frankfurter et al. (1977, lobule VII), for the rat by Rubertone et al. (1995) and for the rabbit by Thunnissen et al. (1989), Epema et al. (1990) and Tan and Gerrits (1992). The general conclusions on the distribution of the vestibulocerebellar projection are depicted in Figs 199 and 200, taken from the papers on the rabbit from the last three authors. Both primary and secondary vestibulocerebellar fibers terminate in lobule X and ventral lobule IX, in the lobule I and II and in the cortex in the depth of the vermal fissures. This projection is mostly ipsilateral for the fibers of the vestibular root and bilateral for the secondary vestibulocerebellar projection. The secondary vestibulo-cerebellar projection to the hemisphere is restricted to the flocculus and the adjacent cortex of the ventral paraflocculus (Fig. 203). A primary vestibulocerebellar projection to the flocculus is absent in the rabbit (Gerrits et al., 1989). Vestibulocerebellar mossy fibers take their origin from neurons in all vestibular nuclei, with the exception of the Deiters' nucleus and a sparse projection from the magnocellular medial vestibular nucleus (Figs 200 and 201). The distribution of neurons projecting to either flocculus or caudal vermis or to both is rather similar and is bilaterally symmetrical. Most neurons were found in the medial, superior and descending vestibular nuclei in this order. Neurons projecting to lobules IX and X, to the flocculus and to both parts of the cerebellum occur in a ratio of 12:4:1 (Epema et al., 1990). A statistical preference was found for the superior vestibular nucleus for a projection to the contralateral flocculus (Tan and Gerrits, 1992). Widespread projections of the nucleus prepositus hypoglossi and neighbouring perihypoglossal nuclei terminate bilaterally with an ipsilateral predominance in the vermis, the flocculus and the paraflocculus and in the cerebellar nuclei (see McCrea and 285

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Fig. 199. Primary and secondary vestibulocerebellar mossy fiber projections in the rabbit, determined with antegrade axonal transport of [3H]leucine and WGA-HRP. Upper panels: sagittal sections; lower panels: transverse sections through the caudal vermis. K196: ipsilateral distribution of fibers of the vestibular nerve. Gerrits et al., (1989); C2098: bilateral distribution of fibers from the medial vestibular nucleus (MV); K82: bilateral distributions of fibers from the superior vestibular nucleus (SV; Thunnissen et al., 1989). Dense termination in the sagittal sections is indicated with heavy hatching, scattered labelled mossy fiber rosettes with light hatching and dots. Note similarity in the distribution of primary and secondary vestibulocerebellar projections. (

Baker, 1985 and Roste, 1989 for reviews). Barmack et al. (1992b) noticed a particularly strong projection of the nucleus prepositus hypoglossi to the ventral paraflocculus (folium p) in the rabbit (Figs 84 and 201). Some cerebellum-projecting neurons of the nucleus prepositus hypoglossi and the caudal parts of the vestibular nuclear complex contain acetylcholine and/or C R F (see Section 3.10.1. and 6.4.5.). Anatomical classifications of the spinocerebellar tracts are based on their level and nuclei of origin, their decussation within the spinal cord, their position in the lateral funiculus, their entrance route into the cerebellum and their lobular and zonal distribution. These different criteria are not necessarily correlated. Oscarsson (1973) in his comprehensive review of the functional organization of spinocerebellar paths, distinguished the classical dorsal and ventral spinocerebellar tracts and added a third tract, the rostral spinocerebellar tract (Oscarsson and Uddenberg 1964), that originates from the cervical cord. Moreover, Oscarsson (1973) included the cuneocerebellar tract as one of the direct spinocerebellar pathways. The dorsal spinocerebellar and cuneocerebellar tracts are ipsilaterally ascending and terminating pathways that transmit information about external events from the lower and upper extremity respectively. Both contain proprioceptive and exteroceptive components. The ventral spinocerebellar and rostral spinocerebellar tracts are hindlimb and forelimb tracts, that convey information to the cerebellum about interneurons mediating flexor reflex afferents and related motor centers in the spinal cord. These tracts terminate bilaterally in the cerebellum. A fifth direct spinocerebellar pathway arises from the central cervical nucleus (Wiksten, 1979b). Although Oscarsson's criteria were functional rather than anatomical and the great flow of new information on the origin and termination of these tracts had not yet started, his classification is still useful. The dorsal spinocerebellar tract takes its origin from Clarke's column and from a group of neurons in Rexed's (1954) dorsal laminae IV-V! (see Yaginuma and Matsushita, 1987; Matsushita and Hosoya, 1979; Matsushita et al., 1979; and Grant and Xu, 1988, and Xu and Grant, 1994, for complete references on rat and cat). It terminates, mainly ipsilaterally, in nine strips in the vermis, the pars intermedia and the extreme lateral part of the lobules III-V of the anterior lobe (Fig. 206C), bilaterally in lobule VIII of the caudal vermis and in parts of the paramedian lobule. The cuneocerebellar projection takes its origin from the external cuneate nucleus and the internal cuneate and gracile nuclei (Gordon and Seed, 1961). The exteroceptive and proprioceptive components of the cuneocerebellar tract synapse in the internal- and external cuneate nuclei respectively (Cooke et al., 1971). The differential termination of exteroceptive and proprioceptive mossy fibers in the apical and basal part of the folia is known from an electrophysiological analysis of the two components of this tract (Ekerot and Larson, 1972). These proximo-distal differences are reflected to some degree in the antegrade tracer studies with injections of the external and internal cuneate nucleus. The projection of the internal cuneate, moreover, is predominantly uncrossed 287

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Fig. 200. Origin of secondary vestibulocerebellar mossy fibers in the rabbit. A-C. Transverse sections through the vestibular nuclei. D. Dorsal view of a reconstruction of the vestibular nuclei. E-G. Transverse sections with retrogradely labelled cells, from injections of fast blue in the lobules IX and X (dots), of nuclear yellow in the flocculus (open circles) and from both injection sites (astarisks). H. Distribution of cells retrogradely labelled from the flocculus. I. Distribution of cells retrogradely labelled from nodulus and uvula, bc = brachium conjunctivum; CO = cochlear nulei; cr = restiform body; d = dorsal part of group y; dfb = direct fastgiobulbar tract; DV - descending vestibular nucleus; gVII = genu of facial nerve; IN = iinterstitial vestibular nucleus; LV = lateral vestibular nucleus; MV = medial vestibular nucleus; MVc = caudal part of medial vestibular nucleus; MVmc = magnocellular medial vestibular nuleus; MVpc = parvocellular medial vestibular nucleus; nV = spinal root of trigeminal nerve; NVpar = parabrachial cerebellar nucleus; PH = nucleus prepositus hypoglossi; S = nucleus of the solitary tract; SV = superior vestibular nucleus; tu = uncinate tract; v = ventral part of group Y; VI = nucleus of the 6th nerve; X = group X of the vestibular nuclei; Y = group y. Bar = 0,6 ram. Epema et al. (1990). (

and directed to the hemisphere (Fig. 202) (Gerrits et al., 1985a; Jasmin and Courville, 1987a,b). The cuneocerebellar tract is distributed to the anterior lobe and the lobules VI, VIII and the paramedian lobule of the posterior lobe (Grant, 1962). The distribution of cuneocerebellar fibers in the hemisphere is complementary to the distribution of the dorsal spinocerebellar tract; in the vermis both projections overlap. This overlap may be related to the predominantly mediolateral organization of somatotopical projections to the anterior vermis and the existence of rostro-caudal somatotopical gradients in the hemisphere in spino-olivo-cerebellar climbing fiber paths (Oscarsson, 1973). Somatotopy in the dorsal spinocerebellar tract was discussed by Xu and Grant (1988) and for the cuneocerebellar projection by Jasmin and Courville (1987b) and Hummelsheim et al. (1985). The ventral spinocerebellar tract is a composite pathway that contains crossed components from lower sacrococcygeal segments, from lumbar spinal border cells and from different cell groups in the lumbar intermediate zone (Matsushita and Hosoya, 1979; Matsushita et al., 1979; Grant et al., 1982; Matsushita and Ikeda, 1980; Matsushita and Yaginuma, 1989 and Yaginuma and Matsushita, 1989). The lower lumbar and sacrococcygeal component terminates preferentially in the apical part of the rostralmost lobules I and II of the anterior lobe. Spinal border cells project to a more extensive area, including the lobules I-V (Fig. 206D). The terminations of the spinal border cells are mainly ipsilateral to their origin, i.e. the fibers recross in the cerebellar commissure. They are distributed to the apical part of the lobules II-V of the anterior lobe (Yaginuma and Matsushita, 1986, Xu and Grant, 1990). The rostral spinocerebellar tract takes its origin from cell groups in Rexed's (1954) laminae VI, VII and VIII of the intermediate zone and a cell group in lamina V of the dorsal horn. The fibers from lamina VIII cross within the cord, the others ascend in the ipsilateral lateral funiculus (Petras, 1977; Petras and Cummings, 1977; Snyder et al., 1978; Matsushita et al., 1978, 1979; Wiksten and Grant, 1980, 1986). The rostral spinocerebellar tract terminates more dorsally than the dorsal and ventral spinocerebellar tract, in the lobules IV, V of the anterior lobe and in the lobules VI, VIII and the paramedian lobule. Their distribution is bilateral, but mainly ipsilateral. Spinocerebellar fibers from lower cervical segments terminate mainly in the vermis of the simple lobule (Matsushita et al., 1985; Matsushita and Ikeda, 1987). The central cervical-cerebellar projection is crossed, with a bilateral distribution in the lobules of the anterior lobe, the bottom of the primary fissure and lobule VIII (Fig. 206A) (Wiksten, 1979a,b; see Matsushita and Tanami, 1987 for a survey of the literature). The termination of the central cervical fibers overlaps with the secondary vestibu289

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locerebellar projection (Matsushita and Wang, 1987) (Fig. 206B) rather than with other spinocerebellar tracts. An important collateral projection to the cerebellar nuclei takes its origin from the central cervical nucleus (Matsushita and Yaginuma, 1995). The central cervical nucleus itself is a site for convergence of vestibulospinal and propriospinal input from neck muscles (Hirai et al., 1978, 1984; Hirai, 1987). Systematic and complete studies with antegradely transported axonal markers on the 290

The cerebellum." chemoarchitecture and anatomy

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distribution of mossy fibers from the paramedian and lateral reticular nuclei are few. A rough idea of their lobular distribution can be gained from the reviews and the papers on retrograde axonal transport of Somana and Walberg (1978), Dietrichs and Walberg (1979b), Gould (1980), Qvist (1989a) and Ruigrok and Cella (1995). The trigeminocerebellar projection recently was studied and reviewed by Ikeda and Matsushita (1992). The basal pontine nuclei, the nucleus reticularis tegmenti pontis and the adjacent paramedian pontine reticular formation are connected with the cerebellum through the middle cerebellar peduncle. Both the nucleus reticularis tegmenti pontis and the paramedian pontine reticular formation send their fibers through the midline raphe into the pes pontis (fibrae rectae), where they deflect laterally to occupy the deep stratum of the middle cerebellar peduncle. The paramedian pontine reticular formation of the cat projects bilaterally to lobule VII and the caudal part of lobule VI and to the ansiform lobule, i.e. to the visual areas of the cerebellum involved in control of saccades (Gerrits and Voogd, 1986; Yamada and Noda, 1987). Fibers of the reticular nucleus of the pons distribute bilaterally, with ipsilateral predominance, to all lobules of the cerebellum, with the exception of the lobules I and X and the dorsal paraflocculus (Kawamura and Hashikawa, 1981; Gerrits and Voogd, 1986). This projection includes the flocculus and the adjacent part of the ventral paraflocculus (Fig. 203) (Gerrits and Voogd, 1989) and collateral projections to the cerebellar nuclei (see Section 5.6.). The projection through the middle cerebellar peduncle of rostral and caudal parts of the pes pontis to the cerebellum is reversed (Von Bechterew, 1885; Spitzer and Karplus, 1907; Voogd, 1964; Voogd et al., 1990). Fibers originating in the ventral and superficial layers of the pes pontis travel in superficial layers of the middle cerebellar peduncle and terminate preferentially in caudal and ventrolateral parts of the cerebellum. Von Bechterew (1885) was able to distinguish this pathway (his 'cerebral', i.e. rostral system) because it acquires its myelin rather late. One of its main constituent is the corticopontocerebellar projection from the visual cortex to the caudal vermis and the paraflocculus. Fibers from caudal and central portions of the pes pontis occupy deeper layers of the peduncle and distribute to more rostral parts of the cerebellum. This pathway corresponds to the early myelinating 'spinal' (i.e. caudal) component of the middle cerebellar peduncle of Von Bechterew (1885). It conveys the cortico-pontocerebellar projection to the anterior lobe. Attempts to analyse the pontocerebellar projection in more detail have shown that the organization of this pathway is extremely complicated. Injections of retrograde tracers in single lobules usually result in bilateral labelling of multiple cell columns or shells. The afferent projections from the cerebral cortex and other sources showed the same degree of dispersion. Our knowledge of informationflow in different cortico-pontocerebellar subsystems, therefore, remains incomplete. The cortical and non-cortical afferents and the efferent connections of the pons were studied and reviewed by Brodal and Jansen (1946), Mower et al. (1979), Rosina and Provini (1980, 1984), R Brodal (1968a,b, 1971, 1972, 1978a,b, 1982, 1987), Gerrits and Voogd (1986, 1987, 1989), Keizer et al. (1984), Ugolini and Kuypers (1986), Gerrits and Voogd (1989), Aas (1989), Bjaalie et al. (1991), R Brodal and Bjaalie (1987, 1992) and Nikundiwe et al. (1994) for the cat; Eisenman (1980), Eisenman and Noback (1980), Azizi et al. (1981, 1985), Anderson and Flumerfelt (1984), Angaut et al. (1985), Kosinski et al. (1986), Leggen et al. (1989), Wells et al. (1989), Mihailoff et al. (1989), Mihailoff (1993), Voogd (1995) and Ruigrok and Cella (1995) for the rat; Mihailoff et al. (1980) for the opossum; R Brodal (1980, 1982), Langer et al. (1985a), Schmahmann

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and Pandya (1989, 1993), Fries (1990), Glickstein et al. (1985, 1994), Stein and Glickstein (1992) and Yhielert and Thier (1993) for the monkey. 6.4.2. Zonal arrangement in the termination of mossy fibers: Correlations with cytochemical maps

During their course as semicircular fibers mossy fibers branch extensively. Some branches are distributed to the anterior and the posterior lobe. This type of branching, which was studied by Heckroth and Eisenman (1988), may explain the mirrored somatotopy which is present in the two lobes. The branches in the medullary rays are of a small calibre. In the granular layer the mossy fibers branch preferentially in a direction across the long axis of the folium (Scheibel, 1977), i.e. with the same orientation as the climbing fibers (Fig. 6). The termination of entire mossy fiber systems in the granular layer often consist of a number of parallel longitudinal arrays of mossy fiber rosettes (van Rossum, 1969). These strips are not as sharply delimited as the terminal zones of the climbing fibers in the molecular layer. Discrete strips often are visible only in the periphery of the projection field; in the center the strips coalesce into a single field. Corresponding concentrations of stem fibers are present in the cerebellar white matter. The termination of mossy fibers in longitudinal strips was first observed and illustrated for the termination of the spinocerebellar fibers in the rabbit (Voogd, 1967; Van Rossum, 1969). It was also reported for the spinocerebellar projections in Tupaia glis and the ferret (Voogd, 1969), the cat (Voogd, 1969 and the series of papers of Matsushita c.s., cited above), Trichosuris vulpecula (Watson et al., 1976), the Virginia opossum (Hazzlet et al., 1971) and the rat (Gravel and Hawkes, 1990; Tolbert et al., 1993). A zonal pattern is also characteristic for the termination of the cuneocerebellar tract in the cat (Voogd, 1969; Gerrits et al., 1985b; Jasmin and Courville, 1987a and b) and rat (Ji and Hawkes, 1994), projection of the lateral reticular nucleus both in cat (Ktinzle, 1975; Russchen, 1976) and rat (Chan-Palay et al., 1977) and in the secondary vestibulocerebellar projections to lobule IX and the anterior lobe (Epema et al., 1985; Matsushita and Wang, 1987) (Fig. 206B). Zonation has not been observed in the distribution of the primary and secondary vestibulocerebellar root fibers to lobule X and the flocculus and was only observed in some parts of the vermis for the projections from the basal pontine nuclei and the reticular nucleus of the pons in the cat and the tree shrew (Voogd, 1969; Gerrits and Voogd, 1986; Kawamura and Hashikawa, 1981). The zonal distribution of spinocerebellar and pontocerebellar fibers in Tupaia glis (Voogd, 1969) is illustrated in Figs 204 and 205, made from silver impregnated sections with large lesions of the pes pontis and low cervical cordotomies. The zonal pattern is striking in the tree shrew and the zonation in the spinocerebellar projection is generally similar to that in other species. A precise comparison of the spinocerebellar and ponto293

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Fig. 203. Mossy fiber projections to the flocculus and the adjacent paraflocculus in the cat. Based on antegrade tracing experiments with tritiated leucine. Notice the lack of basal pontine and reticulopontine projections to the flocculus and their presence in the medial extension (ME) and the caudal lobules (PFLVc) of the ventral paraflocculus in the upper three diagrams. Vestibulo-cerebellar fibers in lower two diagrams terminate both in the flocculus and the ME. A, AP - stereotactic planes; DV = descending vestibular nucleus; FL = flocculus; LV = lateral vestibular nucleus; ME = medial extension of the ventral paraflocculus; MV = medial vestibular nucleus; N P - nuclei pontis = NRTP = nucleus reticularis tegmenti pontis; P F L D - - dorsal paraflocculus; PFLV(c) = (caudal folium of the) ventral paraflocculus; SV - superior vestibular nucleus. Gerrits and Voogd (1989). (

cerebellar projections is not possible, because the experiments were done in different animals. It is not known whether they interdigitate or overlap. More detailed observations on the zonation in the spinocerebellar and trigeminocerebellar projection were reported by Matsushita c.s. for different spinocerebellar pathways in the cat. They plotted the mossy fiber rosettes on reconstructions of the surface of the individual folia. Data on the distribution of mossy fibers on the dorsal (caudal) surface of lobule IV are assembled in Fig. 206. Spinocerebellar fibers from the central cervical nucleus (Fig. 206A) (Matsushita and Tanami, 1987) and the medial vestibular nucleus (Fig. 206B) (Matsushita and Wang, 1987) terminate in three, presumably overlapping zones 1-3 in the bottom of the lobule. Zone 3 is stated to be located at the border of the zones A and B. This interpretation is supported by our map of the distribution of AChE in rostral lobule IV (Fig. 206G). Spinocerebellar fibers from the cervical enlargement are located in the same three zones, in more apical parts of the lobule (Matsushita et al., 1985; Matsushita and Ikeda, 1987). Spinocerebellar fibers from the thoracic cord distribute to the midline and to three parasagittal zones, numbered 2-4. Zone 4 is situated in the medial B zone, a number of patches are present in the lower portion of the hemisphere (Fig. 206C) (Yaginuma and Matsushita, 1987). Spinal border cells project to more apical parts of the hemisphere, with the most medial zone 1 being located at the border of the B and C1 zone (Fig. 206D) (Yaginuma and Matsushita, 1986; Matsushita and Yaginuma, 1989). Fibers from the lower lumbar cord distributed widely over vermis and hemisphere in a distinct zonal pattern (Yaginuma and Matsushita, 1989). The termination of lower lumbar, sacral and coccygeal fibers in the apical parts of the lobules is mostly restricted to a single band around two mm. from the midline, i.e. overlapping the B zone (Matsushita, 1988). Similar plots from the cuneocerebellar (Fig. 206E) and the basal pontocerebellar projection (Fig. 206F) were reproduced from Gerrits (1985). Cuneocerebellar fibers terminate in bands in the vermis and the hemisphere, the gap that separates them presumably corresponds to the B zone. Pontocerebellar fibers are mainly restricted to the apical hemisphere, i.e. the C and D zones, and are scarce in the vermis. The mossy fibers in lobule IV, therefore, belong to different pathways, that terminate in transversely oriented projection fields with a different baso-apical distribution. Vestibulo-, spino- and cuneocerebellar fields can be subdivided into rostro-caudally oriented concentrations of mossy fiber rosettes. The different fields and their concentrations of terminals partially overlap. Correlations between the terminations of mossy fibers and cytochemical maps are rare. Gerrits et al. (1985b) mapped the localization of cuneocerebellar fibers with respect to the borders of white matter compartments in adjacent, H~iggqvist-stained sections (Fig. 202). The projections of low thoracic-lumbar cord to the cerebellum of the rat were compared to the localizations of Zebrin l-immunoreactive Purkinje cells in surface maps of all relevant cerebellar lobules by Gravel and Hawkes (1990). These authors noticed 295

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Fig. 204. Diagrams of the distribution of degenerated, silver impregnated spinocerebellar and pontocerebellar fibers after lesions of the cervical cord and the pes pontis with the nucleus reticularis tegmenti pontis in sagittal (upper panels), transverse (middle panels) and horizontal sections (lower panels) through the cerebellum of Tupaia glis. Note zonal distribution in the vermis and pars intermedia and complementarity of the two projections to the cortex and to the cerebellar nuclei illustrated in middle and lower panels. ANS = antiform lobule; cr = restiform body; fl = primary fissure; FLO = flocculus; ia = anterior interposed nucleus; i p posterior interposed nucleus; L = lateral cerebellar nucleus; m = medial cerebellar nucleus ; PFL = paraflocculus; SI = simple lobule; I-X = lobules I-X. Voogd, unpublished.

a detailed correspondence between the zonal distribution of spinocerebellar fibers in the rat with that reported for other species. The P1 + Zebrin band in the anterior lobe usually overlaps with a spinocerebellar cluster, a second cluster is located under P 1-, few terminals underlie P2+ and a third concentration of mossy fiber rosettes coincides with 296

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P 2 - (i.e. the B zone) extending into P3+ (Fig. 207). Matsushita et al. (1991), who mapped fibers from the cervical cord in Zebrin-I stained sections of rat cerebellum found less correspondence of the concentrations of rosettes with the borders of the immunoreactive Purkinje cell zones. Ji and Hawkes (1994) showed that cuneocerebellar mossy fiber terminals are located between the concentrations of lumbar spinocerebellar mossy fiber rosettes in P1 +, P1- and P2- of lobules II and III of the rat cerebellum (Fig. 207). A close correspondence between multiple patches of mossy fibers with vibrissal receptive fields and the Zebrin-negative P 1-, P2- and P3- zones of lobule IX of the rat cerebellum, was observed by Chockkan and Hawkes (1994). An organization of the spinocerebellar projection in medio-laterally oriented bands, located mainly near the junctions of the lobules I-II, II-III and III-IV (i.e. in the bottom 297

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of the fissures), separated by terminal-free areas and the presence of anterior-posteriorly aligned regions within these bands with high and lower densities of the terminals, was shown in computer reconstructions of the unfolded cerebellar cortex of the rat by Tolbert et al. (1993). It was implied by these authors that the anterior-posteriorly oriented zonation in the spinocerebellar projection was less distinct than had been suggested by previous authors. We would agree that some of the published diagrams of the zonation in the spinocerebellar projection exaggarate the sharpness of this projection. In reality the localization is more diffuse, especially in the bottom of the fissures. Parasagittal focussing in mossy fiber systems is most pronounced in the white matter, in the granular layer the mossy fibers disperse.

6.4.3. The somatotopical organization in mossy fiber pathways The somatotopical organization and the convergence in somatosensory mossy fiber paths has been investigated in great detail in the micromapping studies in the rat and other species using natural stimulation of Welker and his collaborators (see Welker, 1987 299

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for a review). They observed a mosaic of small (0.1-0.3 m m 2) columnar modules or patches in the granular layer that receive short latency somatosensory projections from receptive fields located within a specific body structure (Fig. 208). Receptive fields from specific body structures often are represented in multiple patches on different folia and this distribution is not necessarily longitudinal. Topographical continuities in the periphery are not maintained in the projection to the patches in the granular layer. The arrangement of adjacent patches is asomatotopic and the mossy fiber sources of adjacent patches are disjunctive. This type of fractured somatotopy is found in parts of the hemisphere (Crus I and II, paramedian lobule) and vermis (lobules VIII and IX) in rat, opossum and cat; it is folia-specific and highly reproducible. Branching was found of axon collaterals from single cells in the spinal trigeminal nucleus between pairs of patches with the same receptive fields located on the same or on a different folium, or between patches located in vermis and hemisphere (Woolston et al., 1981). Projections from the somatosensory cortex and the superior colliculus, moreover, were found to conform to the patchy mosaic of projections of the periphery to the granular layer of the rat cerebellar hemisphere (Bower et al., 1981; Kassel, 1980). Welker's observations raise interesting questions about the sites of convergence of mossy fiber pathways mediating peripheral, cortical and tectal information, about the significance of a detailed somatotopical localization (fractured, or longitudinal) in the first link of a mossy fiber-parallel fiber pathway, which would get blurred or even lost in the transvere projections of the parallel fibers and, finally, about the relationship between fractured somatotopy and longitudinal zonation in corticonuclear and climbing fiber pathways and even in the mossy fiber systems themselves. Convergence in mossy fiber pathways may occur at precerebellar level. Convergence of tecto- and corticocerebellar pathways takes place in the pontine nuclei; a direct tectocerebellar pathway does not exist. It is not clear how much of the convergence between peripheral and cortical pathways takes place at a precerebellar level. For certain climbing fiber pathways from the cerebral cortex and the spinal cord it has been shown that this convergence occurs in the sensory relay nuclei (Andersson, 1984). The terminations of certain spinocerebellar tracts certainly overlap with the pontocerebellar projection in the apex and in the hemispheral portions of the lobules (Figs 205 and 206) and the same probably holds for the trigeminocerebellar projection (Jasmin and Courville, 1987a). Systematic convergence of peripheral and cortico-pontine input on the same glomeruli or on the same group of granule cells, therefore, is possible, but has never been studied at the necessary level of precision with anatomical methods. Electrophysiological investigations of somatotopic localization in the granular layer, be it longitudinal zonal (Ekerot and Larson, 1980) or fractured (Bower and Woolston, 1983) both adduced evidence that the same somatotopical pattern is transmitted to the overlying Purkinje cells. These authors found no evidence for lateral spread along the parallel fibers. Llinas (1982) (see also Welker, 1987) explained the preferential connection of granule cells with the Purkinje cells overlying them by the greater greater number of parallel-Purkinje cell synapses on the ascending part of the parallel fiber, but this leaves the function of the long sidebranches of the parallel fiber unexplained. The presence of multiple representations of the same body part in the granular layer is in accordance with the description of mossy fibers as mainly transversely oriented, bilaterally distributed semicircular fibers, which give off collaterals during their course. Multiple representations of the same body part are also present as microzones in different longitudinally oriented climbing fiber systems. Cerebral and peripheral input has been found to converge on to these microzones (Andersson and Eriksson, 1981; 301

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Andersson and Nyqvist, 1983) (see Section 6.3.3.1., Fig. 176). Since zonal borders were never determined in Welker's studies, it is not known whether the mossy fiber patches and the climbing fiber-microzones correspond. A crude, somatotopically arranged convergence of certain climbing and mossy fiber paths from the peripheral nerves and the somatomotor cortex to the cerebellum has been noticed by several authors (Provini et al., 1967, 1968; Ekerot and Larson, 1973, 1980). A topographical correspondence also has been noticed for mossy and climbing fibers containing the same peptides (see next Section 6.3.4.). The convergence of mossy and climbing fibers, that share the same somatotopical and neurochemical properties, on to the same regions of the cerebellar cortex, is remarkable because they reach their targets by quite different routes. This convergence may be explained by the presence of a common developmental factor that determines the distribution of these afferents over the cortex (see Section 6.2.). 6.4.4. Collateral projections of mossy fiber systems to the cerebellar nuclei. The nuclear projection of the red nucleus

Projections from mossy fiber systems to the cerebellar nuclei (see also Section 5.6.) were traced from the spinal cord, the lateral reticular nucleus and the nucleus reticularis tegmenti pontis and adjacent parts of the pontine nuclei. Projections from the vestibular nuclei are still disputed. No collateral projections to the nuclei were traced from the cuneocerebellar system. A special position is taken by the projection from the red nucleus to the interposed nucleus. Evidence for a collateral origin from mossy fibers terminating in the cerebellar cortex was provided by Qvist (1989a and b) with retrograde double-labelling from the cortex and the cerebellar nuclei and by Shinoda et al. (1992) with intraaxonal labelling for the collateral projections from the lateral reticular nucleus, the reticulo-tegmental nucleus and the pontine nuclei of the cat. Spinal projections to the cerebellar nuclei have been reported by several authors in cat (Szentagothai in Eccles et al. 1967; Voogd, 1969; Matsushita and Ikeda, 1970; Ikeda and Matsushita, 1973; Robertson et al., 1983), from the cervical enlargement and the central cervical nucleus in the rat (Matsushita and Yaginuma, 1990, 1995) and Tupaia (Fig.204). They terminate bilaterally in both interposed nuclei and in the fastigial nucleus. The projection from the central cervical nucleus in the rat terminates mainly contralaterally in the rostral fastigial nucleus and mainly in central portions of the anterior and posterior interposed nuclei, excluding the caudal fastigial nucleus, the dorsolateral protuberance, the dorsolateral hump, the lateral cerebellar nucleus and the nucleus of Deiters. The collateral projections from the lateral reticular nucleus terminate in the same regions of the cerebellar nuclei as the direct spinocerebellar projections. According to Matsushita and Ikeda (1976) they are absent from the lateral cerebellar nucleus in the cat, but according to Dietrichs (1983b) certain parts of the lateral nucleus receive lateral reticular afferents. A weak projection of the lateral reticular nucleus to the lateral vestibular nucleus that was described by Dietrichs and Walberg (1979a) in the cat, recently was confirmed in the rat (Ruigrok et al., 1995). Collateral projections from the pontine nuclei were mostly traced from the nucleus reticularis tegmenti pontis. Smaller contributions from the dorsolateral and medial pontine nuclei were found by Gerrits and Voogd (1987) in the cat and Mihailoff (1993) in the rat. Their termination is mostly in the lateral part of the posterior interposed nucleus and in the lateral cerebellar nucleus. The caudal pole of the fastigial nucleus receives a projection in cat and Tupaia (Fig.204). It appears as though the collateral 302

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projections of spinocerebellar and reticulotegmental pontine mossy fibers are found in parts of the nuclei that receive a projection of the Iobules that are strongly innervated by these fibers. This topic was illustrated and discussed by Gerrits and Voogd (1987) for the connections of the nucleus reticularis tegmenti pontis with the cerebellar nuclei and the paraflocculus of the cat. The complementarity in the projections of the spinal cord and the pontine nuclei (including the nucleus reticularis tegmenti pontis) in Tupaia is illustrated in Fig. 204. A connection from the red nucleus to the cerebellar nuclei of the cat was described with the retrograde cell degeneration method by Brodal and Gogstad (1954). This connection was confirmed with retrograde tracers in the rat (Huisman et al., 1983; Naus et al., 1985). Huisman et al. (1983) using fluorescent double-labelling techniques, showed that the rubrocerebellar fibers arise as collaterals from the rubrospinal tract. Rubrocerebellar fibers were found to terminate in the anterior interposed nucleus with axonal degeneration methods in the cat (Courville, 1968; Courville and Brodal, 1966). Walberg and Dietrichs (1986) suggested that the rubronuclear projection is either small or absent. Rubrocerebellar fibers were found to terminate in intermediate parts of the cerebellar cortex (Dietrichs and Walberg, 1983). Retrograde degeneration or labelling of rubral cells in previous studies was explained by the interruption of fibers of passage by large lesions or injections of the cerebellar nuclei. 6.4.5. The chemoarchitecture of mossy fibers It is generally assumed that the distribution of mossy fiber rosettes over the cerebellar cortex is uniform and that all mossy fibers use glutamate as a neurotransmitter (Raymond et al., 1984; Beitz et al., 1986; Clements et al., 1986, 1987; Kaneko et al., 1987, 1989). Intense immunoreactivity, using antibodies against phosphate-activated glutaminase or conjugates of glutamate was present in many neurons of the precerebellar nuclei of the rat, including the basal and reticular tegmental pontine nuclei, the vestibular ganglion, the medial and superior vestibular nuclei and the groups x, y and f, the nucleus prepositus hypoglossi, the external cuneate nucleus, the lateral reticular nucleus and the paramedian reticular formation (Beitz et al., 1986; Kaneko et al., 1989). There is strong evidence, however, for more heterogeneity because certain mossy fibers have been shown to be cholinergic and others to contain several neuroactive peptides. Serotonin has been demonstrated in certain mossy fibers with high resolution autoradiography after topical or ventricular infusion of [3H]serotonin in rodents (Chan-Palay, 1975; Beaudet and Sotelo, 1981). Authors using immunocytochemical methods for serotonin in rat, opossum and cat, however, concluded that varicose serotoninergic fibers distribute throughout the cerebellar cortex, but that few if any terminate as mossy fibers (Bishop and Ho, 1985; Bishop et al., 1985; Takeuchi et al., 1982). The presence of ChAT-immunoreactive mossy fibers in the lobules X and IX of the caudal vermis and the flocculus in several mammalian species was discussed in Section 3.10.1. ChAT-positive mossy fiber rosettes were most numerous in the caudal vermis of the rat, the rosettes were large in X and smaller in ventral lobule IX and the lobules I-III of the anterior lobe (Fig. 83). The ChAT-positive mossy fiber innervation of the flocculus was restricted to the ventral folium and the ventral half of the dorsal folium of this lobule. It was less dense than the innervation of lobule X, but a particularly dense plexus of thin, beaded fibers, which may detach from the mossy fibers, was present in this lobule. ChAT-immunoreactive mossy fibers innervating the lobules I, II, IX and X of the rabbit, were most numerous in the banks of the precentral and posterolateral 303

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fissures. The flocculus receives a sparse innervation, but a heavy concentration of ChAT-positive mossy fibers is present in the ventral paraflocculus of the rabbit. ChAT activity measurements (Fig. 84) revealed that the highest ChAT activity of any cerebellar region in rat, rabbit, cat or monkey ever measured was found for folium 2 of the ventral paraflocculus of the rabbit (Barmack et al., 1992a). The origin of these cholinergic mossy fibers was traced from the caudal medial vestibular nucleus and the nucleus prepositus hypoglossi, with double-labelling of retrogradely transported HRP from injections in the caudal vermis and the flocculus and ChAT immunohistochemistry, both in rat and rabbit (Fig. 201) (Barmack et al., 1992b). Single and double-labelled ChAT-immunoreactive neurons were primarily found in the caudal part of the medial vestibular nucleus, the nucleus prepositus hypoglossi and the vestibular efferent nucleus. They were absent from the superior nucleus. Single HRP labelled cells were present in all vestibular nuclei with the exception of the lateral vestibular nucleus. The cholinergic projection to the ventral paraflocculus of the rabbitis exclusively derived from the nucleus prepositus hypoglossi. In other brain stem nuclei only a few neurons in the lateral reticular nucleus were double labelled for HRP and ChAT following injections in the lobules X and IX in rat and rabbit. Several peptides (enkephalin-ENK, CCK, CRF, calcitonin gene-related peptide, CGRP) have been localized in mossy fibers. When they are present these mossy fibers occur ubiquitously, but they display a preferential localization in certain lobules and, for ENK-CCK and CRF, at least, in midline and parasagittal bands in vermis and paravermis. These concentrations of mossy fiber rosettes usually are aligned with the climbing fiber bands containing the same peptide (Figs 191-195). ENK-, CRF- and CCK-like immunoreactivity in mossy fibers has been reported in the opossum (King et al., 1986a,b, 1987; Cummings and King, 1990; Cummings et al., 1989; King and Bishop, 1990). With respect to their distribution the three peptides differ. Enkephalin containing mossy fibers are limited to the vermis and the the flocculonodular lobe. CRF-containing fibers are more numerous; the vermis of lobule VI and the flocculonodular lobe are densely CRF-innervated. Sagittal and parasagittal concentrations of mossy fibers containing CRF-like immunoreactivity are particularly evident in the anterior lobe and lobules VII-IX. Scattered mossy fibers are present in the hemispheres. CRF and ENK-like immuno-reactivity co-exists in some mossy fibers (and climbing fibers) in the midsagittal and parasagittal bands in the caudal vermis and in the flocculus (Cummings and King, 1990). Numerous CCK-containing mossy fibers are present in all lobules of the opossum cerebellum, with indications of a zonal distribution in the vermis of lobule III and in parts of the caudal vermis (Fig. 193). Enkephalinergic mossy fibers have also been reported in the rat, where they are universally distributed, with a preference for the vermis (Schulman et al., 1981). CRF also occurs in mossy fibers in cat and sheep (Cummings et al., 1988; Cummings, 1989), rat (Van den Dungen et al., 1988) and rabbit (Errico and Barmack, 1993). The distribution of CRF-immunoreactive mossy fibers in the cerebellum of the cat is very similar to the opossum, with concentrations of mossy fibers underlying the stained bands of immunoreactive climbing fibers in vermis and pars intermedia, and heavy labelling in the flocculonodular lobe (Cummings, 1989) (Fig. 194). The localization of CGRP-immunoreactive mossy fibers over the cerebellum of the cat differs substantially from that of the other peptides. They are present in the paraflocculus, the paramedian and ansiform lobules and in the pars intermedia of the simple lobule and the anterior lobe. In the anterior vermis they are located in the apices of the lobules. 304

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In the posterior vermis they are concentrated in the lobules VII and VIII and in the deep part of the granular layer of dorsal lobule IX. They are absent from ventral IX, X and the flocculus. Some CRF-positive mossy fibers can be labelled in monkeys (Saimiri sciureus and Macacafasciculara, Foote and Cha, 1988) but they seem to be less frequent than in other mammalian species. Some of these CRF-immunoreactive rosettes may originate as collaterals from the climbing fibers in these species. Neurons in several precerebellar nuclei which are known to give rise to mossy fibers are immunoreactive for one or more of these peptides. CRF-like immunoreactivity was located in the medial and descending vestibular nuclei (including groups f and x), the nucleus prepositus hypoglossi, the lateral reticular nucleus and other parts of the medullary reticular formation and the solitary tract nucleus in rat (Olschowka et al., 1982; Sakanaka et al., 1987; Van den Dungen et al., 1987, 1988), opossum (Cummings et al., 1989; Cummings and King, 1990) and cat (Cummings, 1989). These authors also found such cells in the external cuneate nucleus of the cat (but not in the opossum) and in the spinal trigeminal nucleus, the locus coeruleus and the raphe nuclei of both species. The profusion of CRF-immunoreactive neurons that could be retrogradely double-labelled from injections in the caudal vermis of the rabbit cerebellum was stressed by Errico and Barmack (1993). Some of these nuclei also contain CCK-like immunoreactive neurons in the opossum and ENK-like immunoreactive cells both in rat and opossum (i.e. the nucleus prepositus hypoglossi, the medial vestibular nucleus, the lateral reticular nucleus, parts of the medullary reticular formation and certain raphe nuclei (Finley et al., 1981; Williams and Dockray, 1983; King et al. 1987; Walker et al., 1988; Cummings and King, 1990). ENK- and CCK immunoreactive neurons could be double-labelled with injections of HRP in the cerebellum of the opossum (Walker et al., 1988; King and Bishop, 1990). CCK/HRP double-labelled neurons also were present in the medial vestibular and prepositus hypoglossi nucleus. Co-existence of these peptides in single neurons of these nuclei has also been demonstrated for ENK with CRF (Cummings and King, 1990) CGRP-containing neurons were double-labelled with HRP from injections in the cerebellum of the cat in the lateral reticular nucleus, the external cuneate nucleus, the descending vestibular nucleus and in the lateral and ventral divisions of the basilar pons (Bishop, 1992). The origin of a major contingent of CGRP-immunoreactive mossy fibers from the pontine nuclei may explain their preferential distribution to the cerebellar hemisphere.

7. P O S T S C R I P T

7.1. BIOCHEMICAL CORRELATES OF CELL TYPES AND FIBER SYSTEMS The chemical neuroanatomy of the cerebellum offers many examples of cell types, grisea and their connections, that can be recognized on the basis of the expression of particular biochemical markers. However, its significance extends far beyond mere recognition of what was already known. In fact, previous unrecognized or neglected cell types, such as the unipolar brush cell (Section 3.6.2.) and the glycinergic interneuron of the cerebellar nuclei (Section 5.3.), and new connections like the multilayer plexus of monoaminergic and cholinergic afferents (Sections 3.8., 3.9. and 3.10.), have been uncovered by virtue of their biochemical properties. Existing cell populations could be split. In 305

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particular, the Purkinje cells can be subdivided into multiple, biochemically distinct subpopulations, distinguished by their contents of certain neuropeptides (Sections 3.1.2. and 3.1.3.) protein kinase C subtypes (PKC~: Section 3.1.5.), receptors (GABAB receptors, muscarinic receptors, and possibly substance P and vasopressin receptors: Sections 3.7.2., 3.10.2. and 6.1.3.) or other biochemical markers (Section 3.1.8.). Several of these biochemically distinct subpopulations are located in longitudinal zones and/or are restricted to certain lobules of the cerebellum. Purkinje cells proved to be particularly rich in markers related to the inositol phosphate-second messenger pathways and the control of intracellular CaZ+-concentration (Section 3.1.4.). Some markers such as cyclic GMPdependent protein kinase, L7 and the InsP3-recptor protein have proven useful to outline the development, the somatodendritic and axonal extent and the ultrastructure of the Purkinje cells (Sections 3.1.4., 3.1.8., 6.2.). Specific markers for granule cells are less numerous. Granule cells, however, uniquely express specific types of glutamate and GABA receptor sububits (see Sections 3.3. and 3.7.). Granule cells are heterogeneous in their expression of calretinin and a zonal distribution has been reported for nitric oxide synthase (Section 3.4.) in granule cells. Among the GABAergic interneurons of the cerebellar cortex, the Golgi cells and the Lugaro cells are biochemically distinct from the basket and stellate cells, in that basket and stellate cells both react with antibodies against calmodulin (Section 3.6.1.), and contain nitric oxide synthase (Section 3.4.). Golgi cells and Lugaro cells both have been identified with type-specific antibodies (Section 3.6.2.). Sub-populations of Golgi cells were distinguished on the basis of their enkephalin- and somatostatin-like immunoreactivities, the co-localization of GABA with glycine-like immunoreactivity (Section 3.6.2.) and the presence of the metabotropic glutamate receptor subunits mGluR5 and mGluR2/3 in (different?) populations of Golgi cells (Section 3.3.2.). Biochemical markers may be helpful in future studies to characterize subtypes of Golgi cells. One factor that hinders such a correlation is our insufficient knowledge of their axonal trajectories of the Golgi cells. Clues for a morphological taxonomy of the Golgi cells are embodied in the finding of a mainly transverse orientation of Golgi cell axons in the direction of the long axis of the folium by De Zeeuw et al. (1994c) and the discovery of the candelabra cell in the cerebellar cortex as a new type of (Golgi?) cell with a transversely oriented axon, distributed to the molecular layer by Lain6 and Axelrad (1994) (Section 2). The distinction of the small, GABAergic neurons of the cerebellar nuclei that give rise to the nucleo-olivary projections, from the large, non-GABAergic relay cells (Mugnaini and Oertel, 1981) (Section 5.2.) certainly represents one of the most consistent correlations between a type of neuron with a specific projection, and its neurotransmitter. However, the dichotomy of the neurons of the cerebellar nuclei into GABAergic and non-GABAergic neurons was complicated by the discovery of small interneurons that co-localize GABA and glycine, by reports on the presence of GABAergic nucleocortical projections (Section 5.3.) and the occurrence of both excitatory aminoacid neurotransmitters as well as acetylcholine, glycine and cholecystokinin in large neurons of the cerebellar nuclei (Section 5.4.). No specific markers are available for mossy and climbing fibers. However, subsets of mossy and climbing fibers can be distinguished by their immunoreactivity towards antibodies against selected neuropeptides (Sections 6.3.4. and 6.4.4.). A subset of secondary vestibulo-cerebellar mossy fibers, taking their origin from the medial and spinal vestibular nuclei and the nucleus prepositus hypoglossi, reacts with antibodies against choline-acetyltransferase (Sections 3.10.1. and 6.4.1 .). 306

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7.2. NEUROTRANSMITTERS AND THEIR RECEPTORS One of the goals of chemical neuroanatomy is to uncover the chemical nature of signalling pathways. As in other parts of the brain the amino acids glutamate and GABA are likely to be the predominant neurotransmitters in the excitatory (mossy, parallel and climbing fiber) and inhibitory (intrinsic cortical and corticonuclear) pathways of the cerebellum respectively (Sections 3.2.1 .-3.2.3.). The characteristics of the excitatory and inhibitory responses of glutamate and GABA in part depend on the type of receptor expressed post-synaptically. Molecular biology has unraveled a great variety of ionotropic and metabotropic glutamate and GABA receptor types, and recent studies have shown that each cell type is provided with a characteristic set of glutamate and GABA receptor types (Tables 2 and 3). In particular granule cells express a large array of glutamate and GABA A receptor subunits. The functional significance of this is puzzling in view of the single type of excitotory mossy fiber and inhibitory Golgi cell input to the granule cells. Studies with transgenic mice may be useful in unravelling the specific roles of particular subunits in the cerebellar circuitry. Diffusely projecting extracerebellar afferents that contain monoamines (noradrenalin, serotonin, histamine and, possibly, dopamine, Sections 3.8. and 5.7.) and acetylcholine (Sections 3.10.1. and 3.10.2.) and their receptors have been identified. They terminate in a multilayer plexus in the cortex and in the cerebellar nuclei. It is generally assumed that these pathways play a modulatory role in the cerebellar circuitry, but their cellular targets are still largely unknown. Their functional importance was highlighted in papers on the cellular physiology of serotonin (e.g. Bishop and Kerr, 1992) and in the studies of the modulation of compensatory eye movements by adrenergic or cholinergic agonists and antagonists by Pompeiano and Collewijn and co-workers (Pompeiano et al., 1991; Tan and Collewijn, 1991, 1992a,b; Tan et al., 1991,1992,1993a,b; Van Neerven et al., 1991; Collewijn et al., 1992). 7.3. LOBULES AND ZONES The stock in trade of the classical neuroanatomist includes the subdivision of the cerebellum in the vestibulocerebellum and the largely somesthetic corpus cerebelli, the modular organization of the cerebellum and the subdivision of the cerebellar nuclei and the zonal and lobular patterns in the termination of mossy and climbing fibers. The distinction of the vestibulocerebellum as a specific subdivision of the cerebellum received support from studies of the transient biochemical properties of its primary vestibular mossy fibers (Morris et al., 1988) (Section 6.4.1.) and its climbing fiber afferents from the dorsal cap (Wassef et al., 1992a,b) (Sections 6.1.5. and 6.2.) during early stages of their development. Early observations on the high context of the vestibulocerebellum of acetylcholinesterase (ACHE) and cholinacetyltransferase (CHAT) were confirmed and extended by the preferential termination of ChAT-immunoreactive mossy fibers in the vestibulocerebellum of different mammalian species (Barmack et al., 1992a,b) (Sections 3.10.1. and 6.4.1.). Calretinin-immunoreactivity differentiates the adult vestibulocerebellum from other lobules by the strong staining of the unipolar brush cells that prevail in the vestibulo-cerebellum, and the relatively low immunoreactivity in granule cells and parallel fibers (Section 6.1.5.). The distribution of calretinin-immunoreactivity also emphasized the extension of the vestibulocerebellum, beyond the posterolateral fissure, into the ventral uvula and the paraflocculus. The compartmental organization of the white matter of the cerebellum as an expres307

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sion of the distribution of the Purkinje cells in longitudinal zones, the precise correspondence in the parasagittal arrangement of the olivocerebellar and the cortico-nuclear projections and the near perfect correlation of the anatomical zones with their electrophysiological counterparts have been known for several decades (Voogd, 1964, 1969; Oscarsson, 1969; Armstrong et al., 1973; Groenewegen and Voogd, 1977; Groenewegen et al., 1979; Voogd and Bigar6, 1980). It was concluded that the output of the cerebellum is organized as a series of modules that incorporate one or more Purkinje cell zones, their cerebellar or vestibular target nuclei, their climbing afferents and the reciprocally organized nucleo-olivary connections. Still, the discovery by Hawkes et al. (1985) (Sections 3.1.8. and 6.1.3.) in the rat, of a longitudinal zonal pattern in the distribution of Zebrinpositive and -negative Purkinje cells, with its perspective of a biochemical diversity of the modules, came as a surprise. As a pattern the zonal distribution of Zebrin was not new; it was preceded by the discovery of the 5'-nucleotidase patttern in the molecular layer of the mouse cerebellum by Scott (1964) (the identity of the two patterns was shown by Eisenman and Hawkes 1989) (see Section 6.1.3.) and of the AChE-positive and -negative zones in the molecular layer of the cerebellum of the cat (Marani and Voogd, 1977) (Sections 3.10.3. and 6.1.1 .). The great impact of the discovery of Zebrin on studies of the cerebellum was due to the unique localization of the Zebrin epitopes in the Purkinje cells, the distinctness of, and the facility in demonstrating the Zebrin pattern in different species with antibodies against Zebrin I and, later, against Zebrin II (Dor6 et al., 1990). The Zebrin pattern is positively or negatively correlated with the distribution of other Purkinje cell markers, discussed by Hawkes (1992) and Leclerc et al. (1992), and in Sections 3.1.8. and 6.1.3. of this chapter. Notably, the distributions of PKC~ (Chen and Hillman, 1993a) (Section 6.1.3.) and of nerve growth factor receptor protein (Section 3.1.10.) in the Purkinje cells conform to the Zebrin pattern. The distribution of certain substances in the Bergmann glia appears to be linked to the Zebrin pattern: this may be true for 5'-nucleotidase (Section 3.5.) and has been established for 3-fucosyl-acetyllactosamine (Bartsch and Mai, 1991) (Section 3.11.) that occurs preferentially in the Bergmann glia of the Zebrin-negative zones. Does the Zebrin pattern result from the interdigitation of two sets of Purkinje cells that differ in their biochemical properties and in their afferent and efferent connections? The truth, probably, is less simple. Purkinje cells of the A and B zones of rat cerebellum, that project to the lateral vestibular nucleus, are uniformly Zebrin-negative and are delimited by Zebrin-positive bands and satellite bands (Fig. 143). Other zones, that can be defined by their corticonuclear and olivocerebellar connections, such as the lateral extension of the A zone of Buisseret-Delmas (1988a), include both Zebrin-positive and Zebrin-negative regions (Fig. 144). Morever, uniformly Zebrin-positive lobules, like lobule VII, the nodulus, the flocculus and the paraflocculus, contain a complex zonal substructure (Sections 6.1.4., 6.1.5. and 6.3.3.3.). The recent maps of the corticonuclear and olivocerebellar connections of the cerebellum of the rat (Buisseret-Delmas and Angaut, 1993) are fairly accurate and based on the same principles as applied in the cat by Groenewegen and Voogd (1977), Groenewegen et al. (1979) and Voogd and Bigar6 (1980). The Figures 145 and 181 that compare the zonal organization of these connections with the Zebrin pattern are still largely hypothetical. There is no doubt, however, that the two patterns are correlated, but there is no simple 1:1 relationship. The borders between corticonuclear and olivocerebellar projection zones often are located within, rather than in between, the Zebrinpositive and -negative zones. The observation that the cerebellar midline divides the 308

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Zebrin-positive P l+ zone in independent left and right subzones (Hawkes and Leclerc, 1987) and the subzonation resulting from the partial overlap of different markers, discussed by Hawkes (1992) and Leclerc et al. (1992) should be taken into account for a more detailed comparison of the two patterns. Do cerebellar modules differ in other respects than in the biochemical properties of their Purkinje cells? True; they differ in the efferent connections of their target nuclei, but their internal organization seems to be very similar. Differences have been reported for the organization of the nucleocortical projections from the different nuclei (Section 5.3.) and it has been claimed that all nucleocortical projections to the C1 and C2 zones of the pars intermedia of the cat take their origin from the posterior interposed nucleus, i.e. from the target nucleus of C2 zone (Trott and Armstrong, 1990). The nucleo-olivary projections from the posterior interposed nucleus to the rostral medial accessory olive is dense (Nelson et al. 1989), and the amount of electrotonic coupling in this olivary subnucleus is the highest in the olive (Llinas and Yarom, 1981). Moreover, a recurrent excitatory loop connects the posterior interposed nucleus, via the Darkschewitsch nucleus and the medial tegmental tract, with the rostral medial acccessory olive (Ruigrok and Voogd, 1995). The C2-nucleus interpositus posterior module, therefore, may be an example of a special type of module, characterized by well-regulated mass action, tight interconnections and a relative lack of internal specialization. The C2 zone, moreover, lacks a somatotopic organization (Section 6.3.3.1.). The C2 module differs in most of these respects from the C1/C3-nucleus interpositus anterior module. Nucleocortical projections from this nucleus are rare or absent (Trott and Armstrong, 1990), the nucleoolivary projection from the anterior interposed nucleus to the dorsal accessory olive (Nelson et al., 1989; Ruigrok and Voogd, 1990) and the degree of electrotonic coupling of the neurons of the subdivision of the inferior olive are rather weak. Moreover, it lacks a recurrent nucleo-mesencephalo-olivary pathway. Both the olivo-cerebellar pathway from the dorsal accessory olive to the C~ and C3 (and d2) zones and the cortico-nuclear projection of these zones to the anterior interposed nucleus display a detailed somatotopical (microzonal) organization (Section 6.3.3.1.). The search for differences in the internal organization of the modules and their biochemical and electrophysiological correlates, therefore, should continue. 7.4. THE ROLE OF BIOCHEMICALLY DEFINED SYSTEMS IN CEREBELLAR MOTOR CONTROL Important contributions of the chemical anatomy of the cerebellum concern our understanding of its role in motor control. The perpendicular arrangement of the parallel fibers and the Purkinje cell zones and microzones is one of the fundamental structural properties of the cerebellum. This arrangement optimalizes the chance that any given mossy fiber input of the cerebellum may interact, through the parallel fibers, with the Purkinje cells or the climbing fibers of a particular zone or microzone. This 'suggests that a mossy fiber input to a restricted part of (a) zone can influence a number of microzones via the parallel fibres. If the strength of the parallel fiber-Purkinje cell transmission is modified separately in each microzone, specific combinations of microzones may be selected for each mossy fiber input. This modification would be performed by the climbing fibers system, which has been shown to exert both short-term (Ebner and Bloedel, 1984) and long-term (Ito et al., 1982; Ekerot and Kano, 1985, 1989) effects on the parallel fiber-Purkinje cell transmission. (...) The propagation of activity along the parallel fibers from (one) zone into the (next) zone would result in output units 309

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involving parts of both zones. Probably spread of activity across the borders of the sagittal zones is a general feature. This would allow far more complex synergies as, for example, combinations of distal and postural muscles' (Garwicz and Andersson, 1992, p. 621). The processes responsible for the selection of the appropriate output for a particular mossy fiber-parallel fiber input may function at the level of the synapses of the mossy fibers with granule cells, unipolar brush cells, or Golgi cells and at the level of the parallel fiber-Purkinje cell synapse. Several systems may be involved in this selection process. Nitric oxide, that is synthetized by subsets of granule cells (Schilling et al., 1994; Hawkes and Turner, 1994) and in stellate and basket cells (Bredt et al., 1990) (Section 3.4.) has been shown to be involved in the production of long-term depression (LTD) of transmission in synapses of the parallel fibers with the Purkinje cell dendritic spines (see Section 3.1.6.), but may also function at the level of the granular layer. Parallel fiber-Purkinje cell transmission also can be blocked by adenosine, acting on the A1adenosine receptors on the parallel fibers (Section 3.5.). The production of adenosine and the levels of the enzyme 5'-nucleotidase in the molecular layer that degrades adenosine monophosphate to adenosine, both are under the influence of the climbing fibers, and, in certain species, 5'-nucleotidase is distributed in bands. An important piece of information on the selection process concerns the heterogeneous distribution at the synapses between the parallel fibers and the Purkinje cell dendritic spines of the ionotropic glutamate receptors, that are blocked during LTD, and of the metabotropic glutamate receptors that elicit this reaction. Somogyi and co-workers have shown that immunoreactivity for the mGluR1 unit of the metabotropic receptor was never associated with the postsynaptic density of the synapse, but was localized at perisynaptic and extrasynaptic sites. This localization is in marked contrast with the ionotropic receptor subunits that are primarily located at the postsynaptic membrane (see Sections 3.3.1. and 3.3.2.). Induction of LTD could be inhibited by in-vitro immunoinactivation of the mGluR1 subunit (Shigemoto et al., 1994) and LTD could not be elicited in transgenic mice that lack this subunit (Aiba et al., 1994). A major drawback of many studies of the chemical neuroanatomy is that they were conducted in only one species, the rat. There is extensive evidence for species differences in the distribution of the synthetizing enzyme of acetylcholine (CHAT), muscarinic cholinergic receptors and acetylcholinesterase (see Section 3.10.), and there is reason to assume that a similar interspecies variability exists for other transmitter systems. The expression of Zebrin by certain subpopulations of Purkinje cells, and the zonal patterns in the distribution of 5'-nucleotidase, only occur in certain species. It is a fortunate coincidence for the experimental neuroscientist that the Zebrin zonal pattern is expressed in rats, but in other species like the cat or macaque monkeys all Purkinje cells are Zebrin-immunoreactive. Many species-differences in the chemical neuroanatomy of the cerebellum may be due to the selectivity of the antibodies employed in the immunocytochemical techniques, but other differences may be real and may reflect true variations in structure or in the transmission and second messenger systems of the cerebellum.

8. ACKNOWLEDGEMENTS The authors wish to express their gratitude to the many scientists and publishers who gave permission to reproduce illustrations from their publication and made the original 310

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prints available to us. Dr. Greet Schalekamp, Dr. Tom Ruigrok and Dr. Doug Hess allowed the use of some of their unpublished material. The secretarial assistance of Edith Klink, the photography of Eddie Dalm and the artwork of Karin Voogd are gratefully acknowledged. Figures 1 and 4 were redrawn from Ramon y Cajal (1911) by Philip Wilson FMAA, AIMI.

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CHAPTER II

The basal ganglia CHARLES R. G E R F E N AND CHARLES J. WILSON

1. INTRODUCTION The basal ganglia are a major neural system which receives cortical inputs, processes these inputs and feeds them back to the cortex via connections through the midbrain and thalamus. While most cortical areas provide inputs to the basal ganglia, including frontal, parietal, temporal and limbic cortices, the thalamic feedback is directed principally to frontal cortical areas, including prefrontal, premotor and supplementary motor areas (Alexander and Crutcher 1990; Alexander et al. 1986, 1990). This thalamic feedback, which parallels ascending cerebellar connections through the thalamus to primary motor cortex (Schell and Strick 1984), involves the basal ganglia in motor function. Diseases of the basal ganglia result in profound movement disorders. However, the complexity and variety of such disorders makes characterizing a typical function that is affected by basal ganglia disorders elusive. In some diseases hypokinetic disorders predominate, such as in Parkinson's disease, whereas hyperkinetic disorders are typical in Huntington's chorea and Tourette's syndrome. Significant advances have been made in recent years that point to specific neuroanatomical and neurochemical substrates involved in these extremes of movement disorders. However, such theories are recognizably simplified and do not explain the full complexity of movement disorders (Albin et al. 1989; DeLong 1990). For example, slowed reaction time that typifies the hypokinetic dysfunction of Parkinson's disease is dependent in part on the context of cues that trigger movements (Brown et al. 1993; Brown and Robbins 1991; Jahanshahi et al. 1993). Further understanding of basal ganglia function incorporates recent work that points to the essential cognitive, motivational and memory components involved in the generation of normal volitional movements. To understand the role that the basal ganglia perform in the complex integration of information involved in the generation of volitional movements its neuroanatomical and neurochemical organization may be broken down into component parts. First, the organization of cortical inputs to the basal ganglia most likely provide the fundamental functional determinants of this neural system. Second, how such cortical inputs are processed is determined by the organization of the subnuclei of the basal ganglia. Included is the organization of the target neurons of cortical inputs in the striatum, connections of these neurons that progress through the basal ganglia, and a variety of local and multisynaptic feedback circuits amongst subnuclei. Third, the organization of the output of the basal ganglia, and the interface with midbrain and thalamic nuclei, determines the effects that are fed back to the frontal cortex. The basal ganglia provide a singular challenge to elucidating functional organization of a neuronal system. Unlike the cortex in which neurons are distributed in distinct Handbook of Chemical Neuroanatomy, Vo112. Integrated Systems of the CNS, Part III L.W. Swanson, A. Bj6rklund and T. H6kfelt, editors 9 1996 Elsevier Science B.V. All rights reserved.

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layers that aid in the characterization of functionally distinct areas, the principal components of the basal ganglia are, for the most part, characterized by a homogeneity of neuronal distribution. However, what the basal ganglia lack in cytoarchitectural features they make up for in the variety of neurochemical markers that are contained in connectionally defined neuronal populations. Modern neuroanatomical methods, which provide detailed mappings of the axonal connections of neurochemically defined neurons have begun to reveal the functional organization of the basal ganglia. The ability to localize a broad spectrum of neurochemical components within identified neurons, aided in recent years by the work of molecular biology that has provided probes for neuroanatomical localization, has had a dual impact. On the one hand, such probes have aided in the characterization of neurochemically defined pathways. On the other hand, the alteration of many of these markers with pharmacologic treatments has aided in understanding how these pathways are functionally interconnected to carry out the processing of cortical inputs by the basal ganglia. Several principles emerge from analysis of the functional organization of the basal ganglia at the systems level (Gerfen 1992). Fundamental to the function of the basal ganglia is the organization of cortical inputs and how the basal ganglia process these inputs. The outputs of the striatum are organized to convert the excitatory inputs from the cortex so as to have antagonistic effects on the output of the basal ganglia. The balance between these antagonistic effects, which determines the output of the basal ganglia, are regulated by the intrinsic circuitry of the striatum and by various feedback loops between the components of the basal ganglia. Some of these feedback loops are regulated by the patch-matrix organization of the striatum, which is the functional equivalent of the laminar organization of the cortex (Gerfen 1989). Finally, the organization of the output of the striatum in relation to the organization of the output of the basal ganglia, which interfaces with the feedback circuits through the thalamus to the frontal cortex, provides the means of extracting certain types of information from the cerebral cortex. We propose that the cortical inputs to the striatum, the major nucleus of the basal ganglia, are a representation of cortico-cortical connections, and that the organization of the basal ganglia reflect this representation.

2. ORGANIZATIONAL OVERVIEW The basal ganglia are composed of a number of subcortical nuclei, which, for the purposes of this review will be regarded as including the striatum, the globus pallidus, the subthalamic nucleus, the entopeduncular nucleus (in cats and rodents) or the internal segment of the globus pallidus (in primates), the substantia nigra and the pedunculopontine nucleus. The principal components of the basal ganglia, as they appear in coronal sections of the rat brain, are diagrammed in Figure 1 and their major connections, in sagittal section, are diagrammed in Figure 2. The striatum, which is composed of caudate, putamen and nucleus accumbens, is the major nucleus of the basal ganglia, in that it is the target of inputs from most areas of the cortex and provides output to the other components of the basal ganglia. Cortical input to the striatum is excitatory (Kitai et al. 1976) with glutamate being the main neurotransmitter used by corticostriatal neurons (Spencer 1976; McGeer et al. 1977). The output of the striatum is inhibitory (Deniau et al. 1976), with all of the output neurons using GABA as a neurotransmitter (Yoshida and Precht 1971). There are two main output streams from the striatum, which have as their common final target GABA 372

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Fig. 1. Diagrams of coronal (frontal) sections through the rat brain at levels from rostral (A) to caudal (F) in which the major components of the basal ganglia are designated. neurons in the entopeduncular nucleus (internal segment of the globus pallidus in primates) and in the substantia nigra pars reticulata (this organization was early described by Nauta and Mehler 1966). One striatal output stream provides a direct path 373

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Fig. 2. Diagrams of sagittal brain sections depicting the major connections of the basal ganglia. A. Descending pathways from the cerebral cortex to the striatum (CP and Acc) and from the striatum through the 'indirect pathway, including the globus pallidus (GP) and subthalamic nuclei (STN) and 'direct pathway' to the output nuclei of the basal ganglia, the entopeduncular nucleus (EP) and substantia nigra pars reticulata (SNr), which provide inputs to the thalamus, superior colliculus (SC) and pedunculopontine nucleus (PPN). B. Feedback pathways within the basal ganglia include 1) thalamo-cortical projections, from the targets of basal ganglia outputs, including the ventral lateral, ventral medial and mediodorsal thalamic nuclei, back to frontal cortical areas; 2) thalamo-striatal projections, from the intralaminar nuclei to the thalamus; and 3) nigro-striatal dopamine (DA) pathway, from midbrain dopamine neurons to the thalamus and frontal cortex. 374

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Fig. 3. Viewof a single striatal spinyprojection neuron, intracellularlyfilledwith biocytin, in a sagittal section of the striatum (A) and at higher magnification (B). Corticofugal fiber fascicles are clearly evident coursing through the striatum. Spinyprojection neurons are labeled within the striatum with calbindin immunoreactivity.

to the entopeduncular nucleus and substantia nigra. The other striatal output stream provides an indirect input to these nuclei, via projections from the striatum to the globus pallidus (external segment of the globus pallidus in primates), which provides an inhibitory input to the subthalamic nucleus, which provides an excitatory input to the GABA neurons of the entopeduncular nucleus and substantia nigra. It is worth emphasizing at the outset that these two output streams have considerable complexity and are not entirely independent. A prime example is the fact that the neurons providing direct projections to the output nuclei of the basal ganglia, also contribute axon collaterals to the indirect pathway. The output nuclei of the basal ganglia are GABA neurons in the entopeduncular nucleus and substantia nigra. These neurons project to ventral tier thalamic nuclei, which project back upon prefrontal and premotor cortical areas. In addition the GABA output neurons of the entopeduncular nucleus project also to the lateral habenula, and those of the substantia nigra project to the superior colliculus and pedunculopontine nuclei. These latter nuclei provide descending projections to motor nuclei, and ascending projections to the thalamus. The GABA output of the basal ganglia provides a tonic inhibition to their projection targets, which is disinhibited by the direct striatal output pathway (Chevalier et al. 1985; Deniau and Chevalier 1985). Thus the major circuit of the basal ganglia is from the cortex, through its component nuclei to thalamic nuclei which project back upon frontal cortical areas. These projections run parallel to cerebellar projections through the thalamus back to motor cortex with the two systems mostly segregated from each other (Schell and Strick 1984). 375

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In addition to these main basal ganglia circuits there are a number of additional connections amongst the components of the basal ganglia that provide short loop feedback systems. One of the major feedback systems is from dopamine neurons in the substantia nigra back to the striatum, the nigrostriatal dopamine system. Other feedback circuits include projections from the globus pallidus to the striatum, from the subthalamic nucleus to the globus pallidus and striatum and from the thalamus back to the striatum. Such feedback circuits contribute significantly to basal ganglia function and their inclusion as additional rather than primary should not be taken as an indication of their importance. 2.1. COMPARISONS BETWEEN RODENTS AND PRIMATES Much of the neuroanatomical work on the basal ganglia that will be described in this review has been carried out in rodents. A reasonable question is whether there are significant differences between the organization of rodents and other animals, notably primates. The most obvious differences between rats and primates are those involving the gross anatomy of the nuclei of the basal ganglia. There are two major examples. The first is the striatum, which in the primate is subdivided by the internal capsule that provides a structural separation between the caudate and putamen nuclei. This structural separation does provide a gross separation of functional regions in the striatum in that the caudate nuclei is the target of prefrontal cortical inputs, whereas the putamen is the target of motor and somatosensory inputs. As the cortical input to the striatum is in large part responsible for its function, the caudate and putamen in the primate are to some extent functionally distinct. However, the internal capsule does not provide a precise divider of functional zones and there is some overlap of inputs from prefrontal cortex to the putamen. In the rodent, which lacks such a distinct structural separation there are nonetheless regional differences in the striatum which are comparable to those of the caudate and putamen, again determined by the regional distribution of inputs from different cortical areas. The second major gross anatomical difference between rats and primates involves the internal segment of the globus pallidus. In primates, this nucleus is situated immediately adjacent to the external segment of the globus pallidus. In rats, the homologous nucleus is separated from the globus pallidus and is embedded in the fiber tract of the internal capsule. In rats, this nucleus is termed the entopeduncular nucleus, which reflects its location. However, the internal segment of the globus pallidus in primates and the entopeduncular nucleus in rats are comparable structures in terms of their connections. Both nuclei represent, along with the substantia nigra pars reticulata, which is nearly identical in both rats and primates, the output structure of the basal ganglia. Despite the gross anatomical differences noted, the major connectional organization of the basal ganglia in rats and primates is remarkably similar. Two of the major features of basal ganglia organization that will be dealt with in some depth in this review, the patch-matrix compartmental organization of the striatum and the organization of direct and indirect output pathways of the striatum, have been demonstrated in both rodents and primates, and appear in the main, nearly identical in organization. Other aspects of the projections of the striatum appear to be also identical, as best demonstrated by papers in which comparable neuroanatomical tract tracing experiments have been done in rats and primates. For example, individual neurons in the striatum have been shown to provide dual inputs to multiple zones in the globus pallidus (Chang et al. 1981; Gerfen et al. 1985; Wilson and Phelan 1982). This dual projection to the globus pallidus has 376

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also been demonstrated to exist in the striatal projection to the substantia nigra in rats (Gerfen et al. 1985). Later, Parent and his co-workers demonstrated a similar organization in primate projections of the striatum (Parent and Hazrati 1993). A second example involves the general topographic organization of the projections of the striatum to the globus pallidus and substantia nigra. In papers that have examined this aspect of striatal organization in the rat (Gerfen 1985) and primate (Parent and Hazrati 1994) using injections of two anterograde tracers into the striatum to chart the topographic distribution of projections to globus pallidus and substantia nigra a remarkably similar pattern of organization is apparent in both. A comparison of the chartings of striatal projections in these two animals from these papers are nearly identical in pattern (Gerfen 1985; Parent and Hazrati 1994). Thus, for the most part, the major organizational principles of basal ganglia organization appear nearly identical in rats and primates. Differences in the organization of the basal ganglia in rats and primates can for the most part be attributed to the expanded cortex in primates. In primates, cortical fields are considerably elaborated and more precisely defined in terms of functional segregation of different cortical areas. While the organization of cortico-striatal patterns appears to follow the same general principles in rodents and primates, the elaboration of more detailed precise mapping patterns appear to predominate in the primate.

3. CEREBRAL CORTEX INPUT TO STRIATUM

The cerebral cortex provides a major input to the striatum. This input originates from most cortical areas, including primary and higher order sensory areas; motor, premotor and prefrontal regions; as well as from limbic cortical areas. It has been well established that this input is organized in a general topographic manner in that the spatial relationships between cortical areas are maintained in the projections to the striatum (Carman et al. 1965; Kemp and Powell 1970; Webster 1961). For example, projections from prefrontal areas are directed mainly to the rostral caudate nucleus (Goldman and Nauta 1977), while cortical inputs from motor cortex terminate primarily in the rostral putamen (Kunzle 1975). More complex is the issue of overlapping projections from functionally related areas. While it is clear that, in general, cortical areas provide input to a much broader area of the striatum than accounted for on the basis of topography alone, the varied and sometimes intricate pattern of this organization have led to a variety of interpretations as to the functional significance. While, the widespread nature of corticostriatal organization is not in doubt, where some have seen patterns of overlap related to patterns of cortical connectivity (Yeterian and Hoesen 1978), others have seen interdigitation (Selemon and Goldman-Rakic 1985). Detailed mapping of the organization of corticostriatal inputs has begun to resolve these issues, showing, in some cases, overlap of inputs from interconnected cortical areas that are organized fairly precisely by the somatotopic organization within such areas (Flaherty and Graybiel 1991; Flaherty and Graybiel 1993a; Malach and Graybiel 1986; Parthasarathy et al. 1992). These issues will be discussed in more detail in a later section. Neurons in layer 5 in most cortical areas provide input to the striatum. All corticostriatal neurons are pyramidal neurons and utilize glutamate as a neurotransmitter. Corticostriatal neurons may be divided into several types based on their connections within the cortex, their projections to other subcortical areas, and their laminar distribution within the cortex. Three corticostriatal cell subtypes have been definitely identified, using double retrograde tracing or intracellular staining or both. Neurons in the frontal 377

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cortex contributing to the pyramidal tract have been shown to contribute axon collaterals to the neostriatum (Cowan and Wilson 1994; Donoghue and Kitai 1981; Landry et al. 1984). The striatal projection of this cell is formed by a very fine collateral formed from the much larger main axon in the course of its trajectory through the internal capsule. While a relatively small component of the corticostriatal pathway, this projection has attracted some interest because of its potential for providing the neostriatum with a copy of the cortical motor signal. A second cortical cell type contributing to the striatum is a bilaterally-projecting corticocortical corticostriatal neuron (Cowan and Wilson 1994; Wilson 1987). This cell is very numerous in agranular cortical regions giving rise to bilateral corticocortical and corticostriatal projections, for example the premotor cortex. Unlike the pyramidal tract neurons, which are located in the deeper part of layer V, these neurons are located in a band at the superficial half of layer V and in the deep half of layer III. The axons of these cells bifurcate twice or more times in the deep cortical layers to form approximately equal-sized branches. Two of them cross the midline to form contralateral projections. An additional branch follows the subcortical white matter laterally to enter the striatum without passing through the striatal part of the internal capsule, and arborizes in the ipsilateral striatum. Additional collaterals travel horizontally, often crossing cytoarchitectonic boundaries to make synaptic connections in other cortical regions on the ipsilateral side. A third corticostriatal neuron has so far been demonstrated only by retrograde labeling (Royce 1983). This is a corticothalamic neuron with a collateral projection to the striatum. It is almost certain that there are other kinds of corticostriatal neurons that have not yet been identified. For example, in cortical regions with few or no collosal projections, such as the granular regions of the somatosensory cortex in the rat, there is a dense band of corticostriatal neurons located in the superficial half of layer V. This band of cells is continuous with the broader band of bilaterally-projecting corticocortical and corticostriatal neurons seen in the motor cortex, but is narrower, and certainly does not have collossal projections. The connections of these cells have not yet been studied in detail. In agranular cortical regions, there are corticostriatal neurons in more superficial layers still, whose identity is unknown. As indicated above, the composition of the corticostriatal projection is greatly dependent upon cortical area, with corticostriatal cells in primary motor and sensory cortices having a much more restricted laminar distribution, and probably a more simple composition, than that of premotor and prefrontal areas (Arikuni and Kubota 1986; Wilson 1987). The intrastriatal axonal arborizations of two corticostriatal cell types in the rat premotor cortex have been described from intracellular staining studies, and were very different, suggesting that each corticostriatal cell type may have a unique pattern of innervation within the striatum (Cowan and Wilson 1994). The striatal collaterals of pyramidal tract neurons made one or more relatively focal arborizations, with dimensions of 100-500 pm on a side. The focal nature of these arborizations suggested a relatively simple topography of the corticostriatal projection formed by these neurons. The other cell type, the bilaterally-projecting corticocortical/corticostriatal neuron, arborized in a much larger striatal volume, with dimensions of 1 mm or greater. Within that volume the axon occupied space in a very sparse fashion, with individual branches running approximately parallel and separated by large uninnervated areas. This pattern is reminiscent of the heterogeneous and complex pattern of labeling seen following small injections of anterograde tracers in the cortex. This pattern is expected from the arborization seen for individual corticostriatal neurons if nearby corticostriatal cells have fine scale similarities in their axonal arborizations. That is, the pattern of labeling seen after 378

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extracellular injections of anterograde tracers in the cortex implies that much of the complex topology of corticostriatal axonal arborizations will be shared among neighboring cells in the cortex. All afferent inputs to the striatum that have been studied so far have formed axonal fields in which the individual axonal branches cross over the dendrites of individual spiny neurons, making synapses mostly en passant. This is the cruciform axodendritic pattern of innervation (Fox et al. 1971), which places each axon into position to contact the maximum number of neurons but minimizes the number of synapses possible with each postsynaptic cell. This is in contrast to the longitudinal axodendritic synaptic arrangement formed by striatopallidal fibers (Fox and Rafols 1976), in which individual axonal branches form multiple synaptic contacts on the dendrites of postsynaptic neurons.

4. STRIATUM The striatum comprises the caudate, putamen and nucleus accumbens. In mammals in which corticofugal fibers coalesce into the internal capsule within the striatum, the caudate nucleus and putamen nucleus are separated by this partition. In animals in which corticofugal fibers are dispersed there is no clear separation between these nuclei, thus the term caudate-putamen is often used. The caudate and putamen, in most species, generally occupy the dorsal part of the striatum. The nucleus accumbens is the rostroventral extension of the striatum, and occupies the area surrounding the anterior commissure in the rostral part of the striatum. The term ventral striatum is generally used to refer to the nucleus accumbens and more caudally, the ventral most part of the striatum (Heimer and Wilson 1975). The olfactory tubercle is sometimes included as a part of the ventral striatum, but in this review will not be discussed. The striatum is composed of one principal neuron cell type, the spiny projection neuron (Bishop et al. 1982; DiFiglia et al. 1976; Wilson and Groves 1980). This spiny projection cell type makes up as much as 95% of the neuron population (Kemp and Powell 1971). These neurons are rather homogeneously distributed such that the striaturn lacks distinct cytoarchitectural organization when all neurons are stained in histologic sections, as contrasted with the laminar organization of the cortex, for example. Using retrograde axonal transport methods Grofova (Grofova 1975) established that these neurons are the projection neuron of the striatum. Cortical input to the striatum targets spiny projection neurons (Somogyi et al. 1981), although not exclusively. Thus the spiny projection neuron is the major input target and the major projection neuron of the striatum. The connections of these neurons are thus the major determinant of the functional organization of the striatum. The remaining striatal neurons are interneurons (Bishop et al. 1982; DiFiglia et al. 1976), in that they do not provide projection axons, but rather distribute axons within the striatum, most of which make synaptic contact with spiny projection neurons. Despite being relatively infrequent, striatal interneurons constitute a variety of morphologic and neurochemically defined types. Among these are the large aspiny neurons, which utilize acetylcholine as a transmitter (Bolam et al. 1984; Kawaguchi and Kubota 1993), and medium aspiny neurons (Bishop et al. 1982; DiFiglia et al. 1976), which utilize GABA as a transmitter (Kita 1993). The latter class of interneurons may be further subdivided on the basis of different peptides and neurochemicals that they contain (Kita 1993; Kubota and Kawaguchi 1993; Kubota et al. 1993). Striatal interneu379

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rons are also rather uniformly distributed within the striatum, although in some cases their axons may be distributed in a heterogeneous manner. 4.1. SPINY PROJECTION N E U R O N Spiny projection neurons take their name from their morphologic appearance (Bishop et al. 1982; Chang et al. 1982; DiFiglia et al. 1976; Wilson and Groves 1980). This neuron cell type is often referred to as the striatal medium spiny neuron. However, as the size discriminator, medium, implies that there are both larger and smaller class neurons, and in the striatum there are no identified smaller neurons, this term is deleted from the classification used in this review and replaced with a more appropriate characteristic feature of these neurons, namely that they are projection neurons. They have a cell body approximately 20-25 pm in diameter, from which radiate 7-10 moderately branched dendrites that are densely laden with spines. The dendrites of an individual neuron extend over an area of approximately 200 pm in diameter. The distribution of the dendrites is not always uniform and may in fact be limited by compartmental boundaries within the striatum, such as those that form the 'patch-matrix' compartments (Kawaguchi et al. 1989). Spiny projection neurons extend a local axon collateral that remains within the striatum. In most cases such collaterals distribute over an area roughly equal in size, but not necessarily in the same area, as the dendrites of the parent neuron (Bishop et al. 1982; Kawaguchi et al. 1990). In some cases the local axon collateral may have an extensive distribution over a very large area within the striatum, extending over 1 mm from the parent neuron (Kawaguchi et al. 1990). Spiny projection neurons also provide an axon collateral which projects out of the striatum to the globus pallidus and/or entopeduncular nucleus/substantia nigra (Kawaguchi et al. 1990). Two major subpopulations of medium spiny neurons, of approximately equal numbers, may be defined on the basis of their projection targets (Beckstead and Cruz 1986; Gerfen and Young 1988; Kawaguchi et al. 1990; Loopuijt and Kooy 1985). One subset, provides an axon projection to the globus pallidus. The other subset provides an axon collateral to the globus pallidus, and additional collaterals to the entopeduncular nucleus and/or the substantia nigra. These projections will be discussed in detail later. Spiny projection neurons all contain glutamic acid decarboxylase (GAD) the synthetic enzyme for the neurotransmitter GABA (Kita and Kitai 1988). In addition, most of those neurons projecting to the globus pallidus alone contain the neuropeptide enkephalin, whereas most of those which project to the substantia nigra contain the neuropeptides substance P and dynorphin (Beckstead and Kersey 1985; Gerfen and Young 1988; Haber and Watson 1983). Spiny projection neurons contain different complements of neurotransmitter receptors, and other proteins that serve to characterize particular subpopulations of striatal output neurons. These will be discussed in further detail below. Spiny projection neurons receive inputs from the cortex, thalamus and amygdala, which make asymmetric synapses on dendritic spines, and to a lesser degree, dendritic shafts. These inputs provide the major excitatory input to these neurons. In addition, a number of inputs from outside the striatum, and from within the striatum provide inputs that function to modify the responsiveness of spiny neurons to the excitatory input. These include inputs from dopamine afferents from the substantia nigra, inhibitory GABA inputs from the axon collaterals of other spiny neurons, inhibitory inputs from GABA (and peptide containing) striatal interneurons, and inputs from cholinergic striatal interneurons. 380

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4.1.1. Cortical input Corticostriatal afferents make synaptic contact primariy with the expanded head of dendritic spines on spiny neurons (Bouyer et al. 1984; Hattori et al. 1978; Kemp and Powell 1970; Somogyi et al. 1981). According to a recent quantitative study in rats (Xu et al. 1989), of all cortical synapses in the striatum, about 90% are formed with dendritic spines, and about 5% with dendritic shafts. The remaining 5% are on somata. While most dendritic spine synapses are certainly formed with spiny projection cells, the smaller number of dendritic and somatic contacts include the combination of all the inputs onto interneurons, as well as contacts made with spiny cell dendritic shafts. The somata receiving cortical inputs generally do not resemble those of spiny neurons. Corticostriatal synapses are almost exclusively asymmetric and contain small rounded vesicles. Although cortical innervation of the striatum is relatively dense, input from any individual corticostriatal axon to an individual striatal spiny neuron is very sparse. Examination of single corticostriatal axonal arborizations suggests that most corticostriatal axons make synapses on a very small proportion of spiny neurons present in the innervated volume, and probably make no more than one synapse on each spiny neuron contacted. This reflects the fact that cortical inputs from individual corticostriatal neurons, is distributed over a relatively large striatal domain and contacts many spiny neurons (Cowan and Wilson 1994). Consistent with the asymmetric character of corticostriatal synapses onto spiny neurons electrophysiologic studies have demonstrated that corticostriatal input evokes a monosynaptic excitatory post-synaptic potential (EPSP) (Kitai et al. 1976; Wilson 1986). At least two types of corticostriatal afferents have been identified, on the basis of the electrophysiologic effects of these inputs (Jinnai and Matsuda 1979; Wilson 1986). One is a fast conducting collateral of neurons projecting to the brainstem and evokes an EPSP with a latency of 3 msec. A second type, which appears to be the major corticostriatal afferent, is a slower conducting afferent that evokes an EPSP with a latency of 10 msec.

4.1.2. Thalamic input Thalamic inputs from from the intralaminar nuclei, including the the parafascicular/ centromedian complex parts, provide inputs to the striatum that are similar to cortical afferents in that they form asymmetric synaptic contacts, and have strong excitatory effects on the spiny cells. Since the pioneering retrograde degeneration studies of (Powell and Cowan 1956) it was believed that the thalamostriatal pathway consisted of a single topographically organized projection tot the neostriatum. More recent studies (Dube et al. 1988; Xu et al. 1991) have shown that this pathway, like the corticostriatal projection, is heterogeneous in nature. There are actually two independent thalamostriatal projections intralaminar nuclear complex, one originating from the parafascicular/ centromedian nuclei and a separate one from rostral parts of the complex including the central lateral and paracentral nuclei. The parafascicular projection, unlike the cortical input, makes its asymmetrical synaptic contacts preferentially with the shafts of dendrites rather than the spines. In one study (Xu et al. 1991), 89% of synapses formed by fibers from the parafascicular nucleus were formed on dendritic shafts in the neostriatum, with only 11% on dendritic spines. This is almost exactly the reverse of the arrangement of cortical axons. The postsynaptic targets of the parafascicular projection have been shown to be spiny neurons, but perhaps neurons of a special class which do 382

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Fig. 6. A-C. Three examples of intracellularly filled striatal spiny projection neurons. Dendrites, shown in black, are densely laden with spines and extend in an area approximately 200-300/lm around the cell body. Local axon collaterals of these neurons, depicted in gray, spread in area approximately 200-400/lm around the cell body, which does not precisely overlap the dendritic spread of these neurons. Adapted from Kawaguchi et al. 1990.

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Fig. 7. The 3 spiny projection neurons from Fig. 6 (A-C) and a fourth spiny projection neuron (D) are drawn to scale in a sagittal section of the striatum showing the denrites (white), local axon collaterals (black) and projection axons of these neurons to the globus pallidus (GP). Spiny projection neurons A and B are of the type that provide a projection axon to the globus pallidus and axon collaterals that extend to the entopeduncular nucleus and/or substantia nigra. Spiny projection neurons C and D provide projection axons that arborize within the globus pallidus but do not extend out of this nucleus. The pallidal axons arborize in two separate zones within the globus pallidus. Spiny projection neuron D is distinguished by the large area over which the axon collateral arborizes within the striatum, extending over 1 cm in areal extent. Also depicted is a large aspiny neuron showing the large area over which its dendrites (white) and axon collaterals (black) extend within the striatum. Adapted from Kawaguchi et al. 1990.

not receive a cortical input or at least do not receive as dense cortical input. In contrast to projections arising from the parafascicular/centromedian nuclei, fibers from the rostral intralaminar nuclei (e.g. central lateral or paracentral nucleus) form synapses similar to those formed by corticostriatal fibers. In the study by Xu et al. (1991), 93% 385

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Fig. 8. A-C) Examples of the axons of type 1 (Kawaguchi et al. 1990) spiny projection neurons in the globus pallidus (GP). Projections of each of these neurons have axons which arborize in the two parts of the globus pallidus, the region immediately adjacent to the striatum and the central region of the globus pallidus. D) The projection axons of examples of type 1 and type 2 spiny projection neurons (Kawaguchi et al., 1990) depicted in a sagittal section. The type 1 neuron is shown to provide a projection axon that terminates within the globus pallidus (GP) and does not extend beyond this nucleus. The type 2 neuron is shown to provide an axon that arborizes within the GP and then extends two collaterals that provide inputs to the entopeduncular nucleus (EP) and substantia nigra, stn: subthalamic nucleus, VTA: ventral tegmental area, SNc: substantia nigra pars

compacta, RR: retrorubral area. Adapted from Kawaguchi et al. 1990.

of these were formed on dendritic spines, and 7% on dendritic shafts. The projections from these two different sets of thalamic nuclei also differ in their innervation of patch and matrix compartments, as described in a subsequent section.

4.1.3. Nigrostriatal dopamine input Inputs from midbrain dopamine neurons that project to the striatum make synaptic contact with spiny neurons. These afferents have been identified at the ultrastructural 386

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level with immunohistochemical localization of either dopamine (Voorn et al. 1986) or the dopamine synethesizing enzyme tyrosine hydroxylase (Arluison et al. 1984; Bouyer et al. 1984; Freund et al. 1984). Most of these afferents make symmetric synapses and contain large round and pleiomorphic vesicles. Of 280 synapses examined by Freund 387

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et al (1984), 59% made synaptic contacts with dendritic spines. Unlike the axospinous synapses formed by cortical or thalamic inputs, these were symmetrical synapses, usually not made on the head of the spine, and these inputs shared the dendritic spine with another bouton forming an asymmetrical synapse (probably from the cerebral cortex or thalamus). Synapses were made onto dendritic shafts in 35% of the cases, and 6% made synapses with somata. It is often suggested that dopaminergic fibers may release dopamine nonsynaptically into the extracellular space, where it could interact with extrasynaptic receptors. Alternatively, dopamine may escape from the synaptic region and diffuse to extrasynaptic receptors on the postsynaptic neuron or other cell processes in the neuropil. With an eye for this possibility, it has been reported that dopaminecontaining synapses are sometimes seen to be in close apposition with the presynaptic part of assymetric forming boutons, presumably of cortical or thalamic origin. However, these close appositions lack synaptic specializations, and they have not been shown to be common than than appositions between any other elements of the neuropil.

4.1.4. Spiny cell local collaterals inputs (GABA and peptide) Spiny projection neurons have axon collaterals within the striatum that make symmetric synaptic contact with other spiny neurons (Wilson and Groves 1980). Axon collaterals of intracellularly labeled spiny neurons were shown to make synaptic contact with the cell soma of spiny neurons (12% of idenified synapses), with the interspine shafts of dendrites (48%) or with the necks of dendritic spines (40%). As with dopamine-containing synapses, collateral axon inputs to the spines contact the spine neck adjacent to asymmetric inputs to the spine heads. Synaptic connections between spiny neurons have also been identified with immunohistochemical markers that are contained in these neurons, including GAD, or either of the peptides substance P or enkephalin. Each of these markers shows a similar synaptic pattern. Each of these markers is contained in different subsets of sources of afferents to spiny neurons, in addition to being contained in the spiny axon collaterals. Thus, in addition to being localized in spiny collaterals GAD is also localized in the axons of some striatal interneurons, and in certain extrinsic afferents such as those from the globus pallidus. Substance P is perhaps a more selective marker for labeling of afferents orgininating from other striatal spiny neurons, although substance P is also localized in some striatal interneurons. Nonetheless, immunohistochemical localization of both GAD (Bolam et al. 1985) and substance P (Bolam and Izzo 1988) in boutons presynaptic to striatal spiny neurons reveal similar distribution patterns. Such inputs are distributed on the cell soma or smooth proximal part of the dendrites, to interspine dendritic shafts or to the dendritic spines. In all cases, the morphological appearance and the distribution of spiny cell collaterals is similar to that of the dopaminergic input. Izzo and Bolam (1986) reported that substance P containing boutons make synaptic contact most often with the more proximal parts of the dendrites, both the soma and smooth parts and the proximal spiny portions. This is somewhat contrasted with the dopamine containing inputs that more frequently target more distal dendritic portions. As will be described in some detail, spiny neurons are subdivided into different subpopulations that are both connectionally and neurochemically distinct, although all share a common morphology and use GABA as a transmitter. Thus, it is of some interest whether the local collaterals of these neurons target neurons of their own subpopulation or those of another subset. Bolam and Izzo (1988) have directly demonstrated that substance P immunoreactive boutons make synaptic contact with striatonigral neurons 388

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(also substance P positive), which establishes that, at least for striatonigral neurons, neurons from the same subpopulations of striatal output neurons make contact with one another. As enkephalin and substance P are contained in different connectionally defined populations of spiny neurons the ultrastuctural localization of synaptic contacts provides some indication of the interactions. In this regard it has been reported that both substance P and enkephalin immunoreactive boutons make synaptic contact with the dendrites of spiny neurons that are immunoreactive negative for the same peptide. This at least raises the possibility that neurons contributing to different output streams are directly contacted by each other, although a more detailed analysis of this question is in order.

4.1.5. Cholinergic input Boutons immunoreactive for choline acetyltransferase (CHAT) make synaptic contacts with striatal spiny neurons as well as other striatal cells (Izzo and Bolam 1988). The cholinergic synapses are symmetric and make contact with the cell somata (20%); dendritic shafts (45%) and with dendritic spines (34%). As with the other symmetrical synapses on dendritic spines, these share the spine with an asymmetrical synapse, usually placed more distally on the spine, similar to afferents from the cerebral cortex and thalamus.

4.1.6. Striatal GABA interneuron inputs In addition to the GABAergic striatal spiny projection neuron, a GABAergic interneuron has been identified within the striatum which comprises approximately 2% of the striatal neuron population. This cell was first positively described using loading with radioactive GABA (Bolam et al. 1983), and was later recognized as a subset of neurons staining more intensely using immunocytochemistry for glutamate decarboxylase (GAD) or GABA (e.g. Bolam et al. 1985). More recently, they have been shown to be positive for the calcium binding protein parvalbumin (Cowan et al. 1990; Gerfen et al. 1985; Kita et al. 1990). These are aspiny interneurons, on average larger than the spiny projection neurons, but smaller than the cholinergic cells. They make numerous symmetrical synapses with the somata and dendrites of spiny neurons, as well as other interneurons. More than any other identified source of input, the synapses from the parvalbumin/GABA interneuron preferentially innervates the somata of spiny neurons (Kita et al. 1990).

4.1.7. Somatostatin interneuron inputs A third type of aspiny striatal interneuron is identified by its immunocytochemical labeling with somatostatin, neuropeptide Y, and NADPH diaphorase. These cells have also been shown to be distinguishable from parvalbumin/GABA interneurons on the basis of morphological and physiological criteria (Kawaguchi 1993). Somatostatin positive synapses are formed mainly on shafts of dendrites and dendritic spines of spiny neurons (Takagi et al. 1983). As before, all the spines involved in this connection receive another asymmetrical synaptic contact. 389

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4.1.8. Other inputs In addition to the dopamine feedback from the substantia nigra, at least two other downstream parts of the basal ganglia provide feedback axons to the striatum. One of these is the globus pallidus, which provides GABAergic input to the striatum (Beckstead 1983; Staines et al. 1981). These have not been studied extensively at the electron microscopic level, however, they may provide a prominent input to the striatum as studies have shown that every pallidal neuron that projects to the substantia nigra provides an axon collateral to the striatum (Staines and Fibiger 1984). In addition, the subthalamic nucleus also provides an input to the striatum. This input is relatively sparse as compared to the density of projections of this nucleus to the substantia nigra and to the globus pallidus (Kita and Kitai 1987). Subthalamic input to the striatum appears to provide asymmetric input to spiny neurons. Although the dopamine input the striatum is the dominant input from the midbrain and brainstem at least two other forebrain projection sytems provide inputs. These include a serotonergic input from the dorsal raphe and a noradrenergic input from the locus coeruleus. Added to the list of sources of afferents to the striatum not covered in depth by this review are those from the amygdala. These inputs will not be dealt with in this review, which does not reflect their probable important contribution to basal ganglia function. 4.2. STRIATAL I N T E R N E U R O N S Striatal interneurons, which extend axons within but not out of the striatum, make up some 5-10% of the striatal neuron population (Bishop et al. 1982; Chang et al. 1982; DiFiglia et al. 1976; Kemp and Powell 1971). This class of neuron presents a variety of morphologically and neurochemically distinct subsets. Two major subtypes are identified on morphologic and neurochemical grounds. One is the neuron type which utilizes acetylcholine as a neurotransmitter, the large aspiny cholinergic striatal neuron (Bolam et al. 1984; Kawaguchi 1992; Kawaguchi 1993; Wilson et al. 1990). The other type, which utilizes GABA as a neurotransmitter, is composed of a number of subtypes, generally termed medium aspiny striatal interneurons (Kita 1993). Striatal neurons, including the interneurons, were once classified according to their somatic diameters. This classification was used primarily because it was compatible with the use of Nissl stains and not because it was sufficient for distinguishing the various cell types. It was successful only insofar as it revealed the presence of a small population of giant cells. Morphological criteria applied to the somatodendritic portion of the cells as they appear after Golgi staining was much more successful, enabling the identification of a number of interneuron types. Again the reason for using somatodendritic morphology in preference to axonal arborizations was based on necessity, rather than choice. The Golgi method did not reliably stain the axon of any of the neurons. However, even using the Golgi method, which was excellent for identification of the spiny cells and the giant aspiny neurons, opinions were divided on the exact number of cell types, suggesting that it was not clearly revealing the identifying characteristics of the interneurons (see, e.g. review in Chang et al. 1982). Nonetheless, several aspiny neuron types could be clearly identified by somatodendritic morphological criteria alone. All authors, beginning with K611iker and his observations on the human striatum (K611iker 1896), have described an aspiny neuron with a large cell body and radiating, sparsely branched dendrites. K611iker reported that this cell had a short axon, but it was 390

The basal ganglia

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described by Ram6n y Cajal (Ramdn y Cajal 1911) as the principal projection neuron of the striatum, and this view was held by most authors until the late 1970's and 1980's when the spiny neuron was shown to be a projection cell and the role of the large cell came into question (e.g. DiFiglia 1976). Subsequent studies employing staining by intracellular dye injection have shown this cell to have a locally-projecting axon, although its axon may arborize over a large distance in the striatum, and may also be myelinated (Bishop et al. 1982; Kawaguchi 1992; Wilson et al. 1990). Some authors distinguish two cell types in this category, one possessing a low density of dendritic spines and fewer dendritic varicosities, and one with smoother, often varicose dendrites (e.g. Chang et al. 1982). This distinction is subtle, and difficult to make when examining any one Golgi-stained neuron, and is justifiably met with a degree of skepticism, but the axons of these two classes of large ceils are also proposed to have dramatically different axonal arborizations. While one is the giant interneuron of K611iker, the other is proposed to be a rare type of projection neuron. Evidence for a large, relatively rare striatal projection neuron has accumulated from retrograde tracing experiments (Grofovfi 1979). Its existence has been confirmed using combined retrograde tracing and Golgi staining (Bolam et al. 1981). In addition, a large striatal neuron can be distinguished in tissue stained for enkephalin immunoreactivity on the basis of its dense investment of terminals positive for that peptide and for GABA (Bolam et al. 1985; Penney et al. 1988). Most large neurons with radiating dendrites, including those shown to be interneurons, show few synapses on the soma and proximal dendrites (Chang et al. 1982; DiFiglia and Carey 1986). A third morphological cell type often has a large soma, and that is the spidery neuron (DiFiglia et al. 1976; Fox et al. 1971; Yelnik et al. 1991). This cell has a very dense dendritic tree that remains near the cell body, with dendrites and an axon that recurves to form a dense network in the region of the soma. This cell is present in a variety of sizes, including some that are among the largest neurons in the striatum (DiFiglia et al. 1976; Yelnik et al. 1991). A much smaller, but otherwise similar version of the cell is also common among the medium and small aspiny cells. Authors disagree on whether the spidery neuron should be considered one type or should be divided in two based on somatic diameter (DiFiglia et al. 1976; Fox et al. 1971; Yelnik et al. 1991). Most have decided to subdivide the spidery cells on the basis of size, but the wide range of somatic diameters seen for cytochemically defined cell types should probably inspire second thoughts (see below). Among the smaller of the aspiny neurons, authors disagree on the number of categories that should be applied, and the criteria offered for distinguishing them are much less convincing. Many of these cells resemble the larger spidery neurons, but others have straighter, less varicose, and less branched dendrites. A quantitative study of the dendritic trees of striatal neurons in the primate (Yelnik et al. 1991) yielded only one clearly distinguishable group of small neurons, a position also taken by Ram6n y Cajal (1911), but most authors have separated the smaller interneurons cells into two groups on the basis of dendritic branching patterns (Chang et al. 1982; DiFiglia et al. 1976). More recently, striatal interneurons have been identified on the basis of immunocytochemical staining for markers known to be associated with neurons that do not have projecting axons. This method of classifying neurons is somewhat less ambiguous than the morphological criteria that were previously applied and the cell classes generated in this way have replaced the morphological ones for most practical purposes. One cell type is the cholinergic interneuron, which on the basis of cell size alone must correspond to one or more of the morphological classes of large neurons (Bolam et al. 1984; Wilson 391

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et al. 1990). A second clear class of interneuron is the one that stains using antibodies for somatostatin, neuropeptide Y, or nitric oxide synthetase (Dawson et al. 1991), all of which primarily stain a single cell population (Pasik et al. 1988; Smith and Parent 1986; Vincent et al. 1983a; Vincent et al. 1983b). A third class of interneurons is identified by its intense staining with a variety of procedures that reveal GABAergic neurons, including radioactive GABA uptake (Bolam et al. 1983), GABA antibodies (Pasik et al. 1988; Smith et al. 1987), and antibodies against glutamic acid decarboxylase (Oertel and Mugnaini 1984; Ribak et al. 1979). This cell was shown in combined GABA uptake and Golgi studies to be an aspiny interneuron (Bolam et al. 1983), and it is clearly distinct from the GABAergic projection neurons, which show a similar but much less intense pattern of staining, and from the somatostatin-containing neurons (Chesselet and Robbins 1989). More recently, this neuron was shown to be specifically stained with antibodies against parvalbumin, a calcium binding protein (Cowan et al. 1990; Gerfen et al. 1985; Kita et al. 1990). These three classes of interneurons differ in average size, but with overlapping distributions so that no one neuron could be identified as belonging to one of the cytochemical classes solely on the basis of its size. In fact, parvalbuminpositive neurons show a very broad range of somatic diameters, and the largest of these cells are in the somatic diameter range of cholinergic neurons (Cowan et al. 1990; Kita et al. 1990; Kubota and Kawaguchi 1993). A more direct comparison of the morphological and cytochemical cell classes requires labeling techniques that allow both cytochemical identification of the neurons and complete enough staining of the dendritic tree for analysis of its subtle features. The combined Golgi stain and 3[H]-GABA uptake study of (Bolam et al. 1983) has provided a description of the GABAergic interneuron, which showed it to be a medium-sized aspiny interneuron. Because interneurons are relatively few, and we have not learned to bias the Golgi method toward preferring any particular cell type, it is difficult to get adequate samples of identified interneurons using this technique. Immunocytochemical studies of the cell show a portion of the dendritic trees of the neurons, and so tempt investigators to make comparisons With cells from Golgi-stained preparations. In general, these have not been convincing, and claims of 'Golgi-like' staining with cell markers in the striatum have usually been somewhat exaggerated. The best approach would be a method that would give both high resolution of the dendritic tree and allow immunocytochemical demonstration of the same cells, and could do this for a significant sample of neurons of each type. This was achieved by Kawaguchi (Kawaguchi 1993) using intracellular injection of biocytin in slices. The cells were visualized using interference optics so that the largest and the smallest neurons could be preferrentially targeted. Because interneurons are enriched at both extremes of somatic size in the neostriatum, this yields a good proportion of interneurons, but does not necessarily represent the population of somatic diameters accurately. Immunocytochemical identification of cells was combined with biocytin using flourescent double-labelling. Using this approach, Kawaguchi (1993) showed that choline acetyltransferase positive neurons have radiating, often sparsely spiny, dendrites and large somata, matching the description of the giant interneuron of K611iker. This conclusion was reinforced by the finding that cells definitely identified as choline acetyltransferase positive exhibited the characteristic physiological properties of giant striatal neurons with long radiating dendrites (Wilson et al. 1990). Parvalbumin-positive interneurons had smaller, denser dendritic fields and intense axonal arborizations very close to the cell of origin. Thus they matched the qualitative appearance of the spidery neurons seen in Golgi studies. Cells positive for NADPH diaphorase (a marker for the somatostatin/NPY cell type) 392

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(black) and axon collateral that arborizes within the striatum (gray). F r o m Wilson et al., 1990. B) Distribution of large aspiny striatal neurons labeled with choline acetyltransferase (CHAT) immunoreactivity in a coronal section of the striatum. The patch compartment was labeled in an adjacent section with calbindin immunoreactivity.

differed from the parvalbumin-positive neurons primarily in the density of their dendritic trees and frequency of branching. These cells had more sparse, radiating dendritic trees that branched less frequently. They also had much more sparse and widespread 393

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axonal arborizations. These results suggest that if the Golgi method had revealed their axons more clearly, these two classes of aspiny interneurons would have been more readily distinguishable from the beginning. There are probably more classes of interneurons that will be revealed as new cytochemical markers become available. For these, as for the ones already discovered, some new approaches to revealing their role in striatal function would be welcome. Because of their relatively small density in the striatum, it must be suspected that they do not function in the traditional roles of interneurons, f~vr example, subserving lateral inhibition in the main information-conveying pathway through the striatum or reciprocal inhibition between opposing groups of striatal projection cells. If they do this, it must be done on a very broad scale, as their numbers are always less than 1/10 that of the spiny neurons on whom they must act. The discovery that some of the neurons release neuromodulators such as somatostatin or nitric oxide, or even acetylcholine (which probably acts primarily as a neuromodulator rather than a neurotransmitter in the striatum) suggests the possibility of spatially global modulatory functions for these cells. Even the parvalbumin neuron, which contains the classical transmitter GABA, seems likely to act in a spatially global fashion, as it has been shown that these cells are meshed into a single network by gap junction interconnections between their dendrites (Kita et al. 1990). 4.2.1. Cholinergic neurons

Striatal neurons which utilize acetyl choline as a neurotransmitter, striatal cholinergic neurons, constitute an important type of interneuron population (Bolam et al. 1984; Kawaguchi 1993; Wilson et al. 1990). As discussed above these neurons have been characterized by morphologic studies due to their large size (Chang et al. 1982; DiFiglia et al. 1976; Kawaguchi 1992; Yelnik et al. 1991), with histochemical staining of acetyl cholinesterase (Fibiger 1982), by immunohistochemical studies employing antibodies directed to the synthetic enzyme choline acetyl transferase (Bolam et al. 1984; Kawaguchi 1993; Wilson et al. 1990), and by intracellular filling studies (Kawaguchi 1992; Wilson et al. 1990). Striatal cholinergic neurons have a very large cell body, up to 40 r in diameter from which extend long aspiny dendrites which may split into secondary and tertiary branches. The dendritic fields may cover an area of over l mm with no apparent orientation in any particular axis. Cholinergic neurons extend an axon, which is both extremely fine but extremely extensive in the area which it covers. The fineness of the axon has made it difficult to identify with immunohistochemical techniques. Intracellular labeling of identified cholinergic neurons has shown axons from individual neurons to extend over an area of as much as 2 mm. Input to striatal cholinergic neurons appears to be derived in the form of both excitatory, asymmetric inputs and symmetric, inhibitory input (Bolam et al. 1984; DiFiglia 1987). Both asymmetric and symmetric synapses are distributed over all portions of the neuron, but appear to be densest on the distal dendrites. Asymmetric input to these neurons resembles that from cortex to the medium spiny neuron in ultrastructure, however, identfication of identified cortical inputs to cholinergic neurons has been elusive. There is electrophysiological evidence of direct monosynaptic input from cortex to cholinergic neurons. At least a portion of the symmetric input to cholinergic neurons contain substance P (Bolam et al. 1986). This input is most likely derived from axons collaterals of medium spiny neurons, specifically from the population of substance P-containing striatonigral neurons. 394

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Fig. 11. A) Diagram of a medium aspiny striatal interneuron that had been intracellularly filled with biocytin. B) Distribution of the two major types of non-cholinergic striatal interneurons, parvalbumin-immunoreactive (IR, black dots) and somatostatin-immunoreactive (IR-white dots), in a coronal section of the striatum relative to the patch compartment. Of note is the greater number ofparvalbumin containing neurons in the dorsolateral striatum compared to the ventromedial striatum and the converse pattern of somatostatin containing neurons. Drawing of medium aspiny neuron provided by H. Kita.

Although it is clear that acetylcholine release is important to striatal function the neuroanatomical substrates by which this is regulated have been difficult to clearly identify. One possible mechanism involves the reported increase in acetylcholine medi395

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ated through activation of substance P receptors. Such a mechanism is supported by anatomical evidence, not only with the demonstration of synaptic contact between substance P-containing boutons and cholinergic neurons (Bolam et al. 1986), but also by the localization of substance P (neurokinin-1) receptor mRNA in cholinergic neurons (Elde et al. 1990; Gerfen 1991).

5. GLOBUS PALLIDUS (external segment) The morphology of the globus pallidus has been well studied at both the light and electron microscopic level (DiFiglia et al. 1982), with a variety of labeling techniques, including Golgi impregnation (Francois et al. 1984; Millhouse 1986; Percheron et al. 1984; Yelnik et al. 1984), immunohistochemistry (DiFiglia et al. 1982) and intracellular labeling (Falls et al. 1982; Kita and Kitai 1994). These studies will be only briefly described in the present paper in the context of issues concerning basal ganglia organization. More thorough treatment of the organization of the globus pallidus may be obtained from the original papers cited above or from a review by DiFiglia (DiFiglia and Rafols 1988). There appear to be two major types of neuron cell types within the globus pallidus (Kita and Kitai 1994). One type, has a moderate to large cell soma from which radiate 3-5 dendrites with secondary and tertiary segments, which are aspinous over their entire length and display some varicosities. The dendrites of these neurons are often long, up to 300-400/lm in length, giving a total maximal dendritic coverage of over 1 mm in some cases. Some aspiny neurons display a discoidal dendritic field in that the dendrites spread mainly in a two dimensional field parallel to the border between the globus pallidus and striatum. Other aspiny neurons have dendrites that cover a volume with a more 3 dimensional distribution. While neurons at the border region between the globus pallidus and striatum often display a discoidal pattern, neurons with discoidal dendritic fields are distributed in the central medial regions of the globus pallidus as well. A second type of globus pallidus neuron is distinguished by the spines distributed on its dendrites. The cell bodies of these neurons are generally smaller than those of the aspiny neurons. However, the size and extent of the dendritic fields appear to be similar for the two types, except that spiny neurons did not display a discoid dendrites. Although all pallidal projection neurons appear to utilize GABA as a transmitter, the differences in morphology are matched with some neurochemical differences. For example, the large discoidal type dendrite bearing neurons contain the calcium bind protein parvalbumin, whereas the other pallidal projection neurons do not (Kita and Kitai 1994). Parvalbumin positive neurons are the more abundant of the two types. The projections of pallidal neurons appear to be somewhat different between the two morphologically and neurochemically distinct pallidal neuron populations (Kita and Kitai 1994; Kita and Kitai 1994). Parvalbumin positive/discoidal dendrite-bearing neurons provide axon collateral projections to the subthalamic nucleus, entopeduncular nucleus and substantia nigra, whereas the descending projection of the parvalbuminnegative pallidal neuron is directed primarily to the subthalamic nucleus. Both neuron types appear to project to the striatum, although not all pallidal neurons provide such a projection. Both types of pallidal neuron provide projections to the subthalamic nucleus, entopeduncular nucleus and substantia nigra and to the striatum. In most cases neurons appear to provide collaterals to the subthalamic nucleus and at least one, and usually, all of the other targets. Neurons in the ventral globus pallidus, which are the 396

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target of the projections from the nucleus accumbens (Groenewegen and Russchen 1984; Haber et al. 1985; Hedreen and DeLong 1991; Swanson and Cowan 1975), have a somewhat different projection profile than more dorsal pallidal neurons (Haber et al. 1985; Haber et al. 1993). In the rat it appears that ventral pallidal neurons provide direct inputs to the mediodorsal thalamus and to the reticular thalamic nucleus (Haber et al. 1985; Mogenson et al. 1987). However, in the primate there appears to be a much sparser or non-existent projection from the ventral pallidum to the mediodorsal thalamus (Haber et al. 1993). Most pallidal neurons may be labeled with GAD immunoreactivity and are thus presumed to utilize GABA as a neurotransmitter (Oertel and Mugnaini 1984; Pasik et al. 1988; Smith et al. 1987). This is consistent with the fact that synaptic contacts of pallidal axon terminals with their target neurons are symmetric (Smith and Bolam 1989, 1990, 1991). In addition to GAD immunopositive neurons in the globus pallidus, there are a scattering of cholinergic neurons within the body of the globus pallidus as well as a large number of cholinergic neurons ventral to the globus pallidus (Fibiger 1982; Grove et al. 1986; Ingham et al. 1985). In as much as these neurons appear to be the target of some projections from both the dorsal and ventral striatum, these cholinergic neurons might be considered to be part of the basal ganglia (Grove et al. 1986). These cholinergic neurons have been shown to provide projections to the cerebral cortex (Fibiger 1982; Grove et al. 1986; Ingham et al. 1985, 1988; Saper 1984). 5.1. SYNAPTIC INPUT Neurons in the globus pallidus receive inputs directly from the striatum (Chang et al. 1981; Hedreen and DeLong 1991; Wilson and Phelan 1982), which are inhibitory (Park et al. 1982) and inputs from the subthalamic nucleus, which are excitatory (Kita and Kitai 1987). Inputs from the striatum appear to be the dominant input to pallidal neurons and display a distinct synaptic organization (DiFiglia et al. 1982). Individual fibers from the striatum entwine dendrites of pallidal neurons, making numerous synaptic contacts along an extended region of a dendrite. These synapses are symmetric and on the order of lctm in diameter. These afferents have been demonstrated to contain both GAD and enkephalin-immunoreactivity, consistent with their origin from the striatum. Some estimates place the percentage of such synapses as over 80% of those within the globus pallidus. The manner in which these afferents entwine dendrites forming a mosaic pattern of large synapses gives the globus pallidus appearance of being comprised of radial fibers (DiFiglia et al. 1982). An additional feature of pallidal architecture which enhances this appearance is the bundling of dendrites of separate neurons (Millhouse 1986). The synaptic organization of the globus pallidus, where afferent axons make multiple contacts thus appearing to ensheath pallidal dendrites concerns the possible consequence on convergence of striatal afferents. The radial orientation of pallidal neuron dendrites, orthogonal to the plane of striatal efferent fibers, had suggested a means of convergence in that individual pallidal neurons would spread dendrites across the paths of outputs of many regions of the striatum. However, an alternative organization is suggested by the fact that individual striatal efferents, rather than remaining 'on course' as they traverse the globus pallidus, in fact follow local paths to entwine individual pallidal neuron dendrites. This might suggest that in fact individual striatal efferent neurons make a rather direct transfer to few rather than many pallidal neurons. Such an organ-

397

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Fig. 12. Diagrams of neurons in the globus pallidus that had been intracellularly filled with biocytin. A and B) Globus pallidus neurons with dendrites distributed in a discoid manner, that is primarily within a single plane. The fiat plane of the discoid distribution is parallel to the border between the striatum and globus pallidus. This type of neuron is distributed throughout the globus pallidus. For the most part dendrites are relatively spine free. These neurons emit an axon collateral that arborizes within the globus pallidus (not shown) and also provides projections out of the nucleus (for neuron A see Fig. 13). C and D) Examples of globus pallidus neurons with dendrites that radiate in all planes around the cell body. These dendrites possess spines. Some of these neurons have axons that arborize extensively within the globus pallidus (shown for neuron D in gray) and axons that project out of the nucleus (not shown). E) An example of a striatal spiny projection neuron drawn to the same scale as the pallidal neurons for comparison. Globus pallidal neurons are redrawn from Kita and Kitai (1994).

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1.0 mm Fig. 13. A) Diagram showing an example of inputs to the globus pallidus (GP) from striatal spiny projection neurons. Typically there are two major sites of axonal arborization, one in the region immediately adjacent to the striatum and a second in the central region of the GR B) Stylized drawing of two pallidal neurons showing how the dendrites of neurons are confined within the two regions of the GP that conform to the pattern of striatal inputs. C) The axonal projection of a globus pallidus neuron of the type with discoid dendrites, which provides collaterals to the striatum (CP), to the entopeduncular nucleus (EP), subthalamic nucleus (stn) and substantia nigra (SN). Adapted from Kita and Kitai 1994.

ization would be decidedly different from that of cortical afferents to the striatum, in which individual axons contact the dendrites of many neurons 'en passant'. A second less frequent type of synapse forms asymmetric synapses along all portions of the dendrites of pallidal neurons. These inputs have been demonstrated, using anterograde tracing with PHA-L, to arise from the subthalamic nucleus (Kita and Kitai 1987). 5.2. OUTPUT Descending output of the globus pallidus to other components of the basal ganglia is directed principally to the subthalamic nucleus and to the entopeduncular nucleus (internal segment of the globus pallidus in primates) and the substantia nigra (Haber 399

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et al. 1985, 1993; Kita and Kitai 1994). Ascending outputs of the globus pallidus provide feedback to the striatum (Beckstead 1983; Staines et al. 1981; Staines and Fibiger 1984). In addition, there is a variable projection from the ventral pallidum to the thalamus (Haber et al. 1985, 1993; Mogenson et al. 1987). The source of efferents to the different targets of the pallidal outputs arises from different morphologically and neurochemically defined neuronal types (Kita and Kitai 1994a,b). Of particular note is the synaptic organization of pallidal projection terminals, particularly those that provide input to the internal pallidal and substantia nigra neurons. Pallidal afferents onto these neurons is directed to the cell soma and proximal dendrites, whereas the striatal afferent input is directed to the same neurons' more distal dendrites (Smith and Bolam 1989, 1990, 1991). The organization of these projections will be discussed in more detail below.

6. SUBTHALAMIC NUCLEUS Based on cellular and dendritic morphology neurons in the subthalamic nucleus appear to be of one main type, which nonetheless show a variance in the dimensions of the cell soma and dendritic ramifications (Kita et al. 1983a; Yelnik and Percheron 1979). In rats, the cell somata ovoid or polygonal with a medium size ranging 11-18 r in diameter. Most subthalamic neurons extend 3-4 primary dendrites which taper and branch into secondary and tertiary dendrites. Dendrites show infrequent spines, which, if present, are located on more distal parts of the dendrites. The dendrites spread in varying patterns within the nucleus. In general dendrites appear to distribute in an ovoid area in both the frontal and sagittal planes, thus showing a greater extension in the rostrocaudal dimension than in the dorsal and ventral dimension. In the horizontal plane, dendrites appear to distribute roughly equally in the medial lateral dimension as in the rostro-caudal dimension. Subthalamic neurons across species appear to be similar in morphologic type, although the planar distribution patterns of the dendrites vary from species to species. This presumably reflects different geometries of the afferent inputs in different species. Neurons in the subthalamic nucleus appear to be of one neurochemical type in that most are immunoreactive for glutamate. This is consistent with the fact thta the synapses of subthalamic afferents to neurons in the globus pallidus, entopeduncular nucleus and substantia nigra are asymmetric (Kita and Kitai 1987). Moreover, the electrophysiologic response of neurons postsynaptic to subthalamic afferents following stimulation of the subthalamic nucleus confirms the excitatory nature of these inputs (Nakanishi et al. 1987b; Robeldo and F6ger 1990) 6.1. SYNAPTIC INPUT Neurons in the subthalamic nucleus receive inputs from the globus pallidus, which are inhibitory (Kita et al. 1983b) and inputs from the cortex, which are excitatory (Kita et al. 1983b; Nakanishi et al. 1987a, 1988). Inputs from the cortex are asymmetric and distributed to principally to the dendrites of the neurons. Inputs from the globus pallidus make large symmetric contact which are directed relatively equally to the cell soma (30%), proximal (39%) or distal (31%) dendrites (Smith et al. 1990). This input is distinguished from pallidal inputs to the substantia nigra in which 90% of the synaptic contact is made with the soma or proximal dendrites (Smith and Bolam 1990). 400

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6.2. OUTPUT Neurons in the subthalamic nucleus project axons that target neurons in the globus pallidus and in the entopeduncular nucleus and substantia nigra, as well as a sparse projection to the striatum (Kita and Kitai 1987). These inputs provide an excitatory input to each of the target structures (Kita and Kitai 1991; Nakanishi et al. 1987b; Robeldo and F6ger 1990).

7. SUBSTANTIA NIGRA/ENTOPEDUNCULAR NUCLEUS Together, the substantia nigra and entopeduncular nucleus (internal segment of the globus pallidus in primates) may be considered output nuclei of the basal ganglia in that they provide the interface with brain areas outside the basal ganglia, in particular the thalamus and midbrain structures including the superior colliculus and pedunculopontine nucleus. The neurons that provide these output projections utilize GABA as a transmitter and form a nuclear complex that is continuous from the entopeduncular nucleus (internal segement of the globus pallidus in primates) and substantia nigra pars reticulata. In addition to the GABA neurons in these nuclei, dopamine neurons in the substantia nigra pars compacta provide a feedback pathway to the striatum. The substantia nigra is composed of two main neuron cell types, those that utilize dopamine (Bj6rklund and Lindvall 1984) and those that utilize GABA as a neurotransmitter (Oertel and Mugnaini 1984; Pasik et al. 1988; Ribak et al. 1979). Dopamine neurons are located primarily in the pars compacta, which is a neuron dense zone forming the dorsal part of the substantia nigra (Gerfen et al. 1987). In addition, dopamine neurons are also located in groupings in the ventral neuron sparse zone, the pars reticulata. GABA neurons are localized, for the most part, in the pars reticulata. Dopamine neurons in the substantia nigra, as well as those in the adjacent ventral tegmental area and retrorubral area provide inputs to the striatum (Beckstead 1979; Gerfen et al. 1987; Oertel and Mugnaini 1984; Pasik et al. 1988; Ribak et al. 1979). GABA neurons in the pars reticulata provide inputs to the thalamus, superior colliculus and pedunculopontine nucleus (Beckstead 1979; Gerfen et al. 1982; Oertel and Mugnaini 1984; Pasik et al. 1988; Ribak et al. 1979). Neurons in the substantia nigra have been difficult to classify on the basis of morphologic criteria as classes that may be distinguished clearly by cell body size, dendritic morphology or dendritic spread do not appear (Grofova et al. 1982; Yelnik et al. 1987). The connectional and neurochemical determinants of the two major types of neurons in the substantia nigra do not relate to distinct differences in morphology, although in primates the dimensions of dopamine-containing pars compacta neurons appear to be on average larger than their pars reticulata counterparts. Thus, a generic substantia nigra neuron might be described, with the realization that specific parts of these neurons display a rather wide range of size and shapes. Neurons in the substantia nigra have a medium to large sized irregularly shaped cell soma with axis dimensions ranging from 16-50/~m (long axis) and 8-32/lm (short axis). Several ( 2 4 ) main dendrites radiate from the cell soma and extend over a generally large domain. Dendrites may divide into secondary or tertiary branches of similar size to the main branches, but in general branching is rather restricted. In some cases much smaller, unbranched processes may issue from larger dendrites. The distribution of the dendritic fields is of particular interest due to the organization 402

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of afferent fibers (Francois et al. 1987). In the rat dendritic fields may extend as much as 1-1.5 mm in the rostro-caudal axis, 700 r mediolaterally, and 400 r in the dorso-ventral axis, which covers as much as 70% of the mediolateral dimension and 50-80% of the length of the nucleus (Grofova et al. 1982). However, the orientation of the dendrites of individual neurons varies dependent on the location of the neuron within the pars reticulata. Neurons in the dorsal part of the nucleus have dendrites that spread in all three axis, while neurons in the ventral part of the nucleus have dendrites that remain confined to the ventral plane of the nucleus. Similar organization of neuronal dendritic patterns have been described in the primate as well. As will be discussed in some detail below the organization of the dendritic distributions of pars reticulata neurons is related to the organization of afferent input from the striatum, globus pallidus and subthalamic nucleus (Gerfen 1985; Kita and Kitai 1987). Dopamine-containing neurons in the substantia nigra are similar in many respects to the morphology of pars reticulata neurons (Tepper et al. 1987). The cell somata appear somewhat larger than those of pars reticulata neurons, although, the morphology of the dendrites appear similar. The dendritic distribution of dopamine neurons reveals two distinct populations of neurons. One population is situated in the dorsal part of the pars compacta and possess dendrites which distribute in the plane of the pars compacta. A second population is situated in the ventral part of the pars compacta and in cell groups in the pars reticulata. These neurons posssess dendrites that extend into the pars reticulata. These two populations are also distinct in their efferent projections and neurochemical content, which will be discussed in terms of striatal patch-matrix compartmental organization below. 7.1. SYNAPTIC INPUT TO PARS RETICULATA NEURONS The major sources of input to substantia nigra neurons are GABA inhibitory inputs from the striatum and globus pallidus, and excitatory inputs from the subthalamic nucleus. That the striatum provides an inhibitory, GABAergic input to pars reticulata neurons has been established using electrophysiologic techniques (Chevalier et al. 1985; Deniau and Chevalier 1985; Deniau et al. 1976). The globus pallidus has more recently been established to provide a similar inhibitory input. The synaptic organization of this and the pallidal input to the pars reticulata was described in a comprehensive analysis by Smith and Bolam (Smith and Bolam 1989, 1990; Smith et al. 1990), axonally transported tracer labeling of striatal and pallidal input to identifed pars reticulata neurons projecting to the superior colliculus were examined at the light and electron microscopic level. Striatal input to pars reticulata neurons form symmetric, relatively small, synapses directed principally to distal parts of the dendrites (77% of such input), and only infrequently to the cell soma (3%). In contrast, inputs from the globus pallidus for symmetric, relatively large, synapses directed principally to the perikarya (54% of such input), or to proximal dendrites (32%). The differential distribution of inputs from the striatum and globus pallidus to the distal and more proximal dendrites suggests that, if the inputs are comparable in number, the latter afferent system may exert a dominant control over these pars reticulata neurons. Inputs from the subthalamic nucleus to the pars reticulata provide an excitatory input mediated by the neurotransmitter glutamate (Kita and Kitai 1987; Nakanishi et al. 1987b). At the synaptic level these inputs form asymmetric contacts principally directed to more distal parts of the dendrites of pars reticulata neurons (Kita and Kitai 1987). Thus the distribution pattern of these afferents is similar to that of the striatal inputs. 403

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SNc dopamine neurons

A 1O0 pm

B

Fig. 15. Drawings of two dopamine neurons in the substantia nigra pars compacta that have been intracellularly filled. A) This example is typical of pars compacta neurons that are in the dorsal tier of the nucleus, whose dendrites remain within the pars compacta, for the most part. B) This neuron is an example of pars compacta neurons in the ventral tier of the nucleus which extend dendrites downward into the pars reticulata. Adapted from Tepper et al. 1987.

7.2. SYNAPTIC INPUT TO PARS COMPACTA NEURONS Input to pars compacta dopamine neurons appears for the most part to be similar to that to the pars reticulata for each of the sources of input described above. Input from the striatum, which is identified both directly with anterograde axonal markers, with GABA or with substance P immunoreactivity, appears to provide a major input to pars compacta neurons (Bolam and Smith 1990). However, in the case of input from the globus pallidus the input is somewhat less than that to the pars reticulata neurons (Smith and Bolam 1990). In addition there are other known sources of inputs directed to the pars compacta, that have not been described as being directed to the pars reticulata. One of these is a cholinergic input which provides asymmetric synaptic contacts with pars compacta neurons. Another is from the amygdala, which appears to provide inputs to the major components of the dopamine cell groups, but not to the pars reticulata (Gonzales and Chesselet 1990). In addition, the lateral habenula provides input directed to the pars compacta (Herkenham and Nauta 1979), which has been identified with electrophysiologic techniques as an inhbitory input (Christoph et al. 1986). 404

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Fig. 16. Immunohistochemical labeling of adjacent sections through the rostral part of the substantia nigra showing tyrosine hydroxylase- (A), calbindin- (B) and parvalbumin-(C) immunoreactivity. A) Dopamine containing neurons labeled with tyrosine hydroxylase are distributed in the ventral tegmental area (VTA) and substantia nigra pars compacta (SNc). Of note are the dendrites of SNc neurons that extend downward into the subtantia nigra pars reticulata (SNr). B) Calbindin immunoreactivity labels dorsal tier dopamine neurons in the ventral tegmental area and dorsal tier of the SNc. Ventral tier dopamine neurons in the SNc are not labeled. Calbindin immunoreactive terminals originating from spiny projection neurons in the striatal matrix compartment are distributed in the pars reticulata (SNr) but not in the pars compacta (SNc). This pattern reflects the fact that calbindin-containing matrix spiny projection neurons provide inputs to the SNr, whereas calbindin-negative patch spiny projection neurons provide inputs to the SNc. C) Parvalbumin-immunoreactive neurons are located in the substantia nigra pars reticulata. These neurons are the GABA-containing neurons that provide projections to the thalamus, superior colliculus and pedunculopontine nucleus. From Gerfen et al. 1985.

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Summary of inputs to Substantiia nigra pars retiiculata neurons

sagittal section

B

1 mm

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input from subthalamic nucleus input from striatum input from GP

Fig. 18. Summary of inputs to substantia nigra pars reticulata neurons. A) Stylized drawing showing distribution of dendrites of neurons in the two main regions of the substantia nigra pars reticulata (SNr). Two subregions of the SNr are depicted, similar to those of the globus pallidus, in that striatal afferents and the dendrites of the target neurons in demarcate these separate regions. The distribution of striatal afferents to these two subregions are diagrammed in Fig. 23. Dopamine neurons are located in the dorsal subtantia nigra pars compacta (SNc) and in islands of dopamine neurons (SNc)that separate the two parts of the SNr. B) Stylized diagram of the major synaptic inputs to substantia nigra pars reticulata neurons. GABA-containing terminals from the striatal spiny projection neurons make synaptic contact with the distal portions of the dendrites, with individual fibers making multiple contacts with individual dendrites. GABA-containing terminals from the globus pallidus make synaptic contact with the cell body and proximal dendrites of pars reticulata neurons. Glutamate-containing terminals from the subthalamic nucleus make synaptic contact with the distal portions of pars reticulata neuron dendrites, and similar to those from the striatum, individual fibers make multiple contacts with individual dendrites.

7.3. P R O J E C T I O N S

OF PARS RETICULATA

NEURONS

O u t p u t t a r g e t s o f the s u b s t a n t i a n i g r a i n c l u d e the following: the t h a l a m u s , s u p e r i o r colliculus a n d the p e d u n c u l o p o n t i n e n u c l e u s ( B e c k s t e a d 1979; D e n i a u a n d C h e v a l i e r 1992; G e r f e n et al. 1982; K i t a a n d K i t a i 1987; N a k a n i s h i et al. 1987b). N i g r a l i n p u t s to the t h a l a m u s are d i r e c t e d to t w o m a i n p a r t s o f the t h a l a m u s . T h e first are the set o f nuclei, i n c l u d i n g the i n t r a l a m i n a r nuclei, w h i c h p r o j e c t b a c k to the s t r i a t u m . T h e s e c o n d

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major output of GABA neurons of entopeduncular nucleus and substantia n_.ii

A (sagittal)

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Ih 9

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PPN

B(coronal)

1.0 mm Fig. 19. Diagram of major output of GABA-containing neurons of entopeduncular nucleus (EP) and substantia nigra pars reticulata (SNr). A) Sagittal section showing with white arrows the projections from EP to the thalamus (ventral lateral, vl and lateral habenula, lh) and with black arrows the projections from the SNr to the thalamus (mediodorsal, md; ventral medial, vm and parafascicular-intralaminar complex, pf, il), to the superior colliculus and pedunculopontine nucleus (PPN). B and C) Coronal sections showing thalamic nuclei innervated by the entopeduncular nucleus (with white stippling) and by the substantia nigra pars reticulata (with black stippling).

thalamic target are nuclei which provide projections to frontal cortical areas. The specific nuclei involved, vary from species to species, primarily as a consequence of the organization of cortex. For example, in rodents, the principal target of the substantia nigra is the ventral medial thalamus, which provides a relatively widespread and distributed input to frontal cortical areas, and to the paralaminar medial dorsal thalamus, which projects to the cortical areas thought to be equivalent to the frontal eye fields. Conversely, in primates where frontal cortical areas are subdivided into more discrete cortical areas, thalamic inputs to these areas are correspondingly organized. In primates, the principal thalamic targets of the internal segment of the globus pallidus are the 408

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ventral lateral, pars oralis and ventral anterior, pars parvocellularis nuclei (Schell and Strick 1984), and the target of the substantia nigra is the ventral anterior (VAmc) and paralaminar medial dorsal (MDpc) nuclei (Ilinsky et al. 1985). Many individual pars reticulata neurons have collaterals that target two or more of these targets. The organization of these outputs will be described in more detail.

8. CONNECTIONAL ORGANIZATION OF BASAL GANGLIA The functional organization of the basal ganglia may be considered by breaking down the components of processing that occur as cortical inputs are transformed through the system. First, we will describe the organization of the cortical inputs to the striatum and the continuation of this organization through the circuits of the basal ganglia. There are several determinants of this organization. On the one hand there is a topographic organization that appears to provide for distinct parallel streams from functionally related cortical areas that are processed through the basal ganglia. On the other hand, within each of these zones there is considerable overlap of inputs from widely dispersed cortical areas. We will suggest that the principal organizing scheme in corticostriatal inputs reflects the mapping of the connections of cortico-cortical connections into the striatum. This organization provides the basis of the information that is processed by the basal ganglia. Second, we will describe the organization of projections from the striatum to the globus pallidus and substantia nigra in terms of their organization with the targets of the basal ganglia outputs, principally the thalamus and midbrain structures including the superior colliculus and pedunculopontine nucleus. The organization of these systems reflects in part the organizing principles related to corticostriatal inputs. A second organization emerges in that at each level of projection of one nucleus of the basal ganglia onto another there are dual projection fields. These appear to be related to the organization of the targets of the output structures, and presumably reflect the nature of the information that is provided by the cortex. Third, we will describe the organization of systems that regulate the dopamine feedback system. This organization relates to the 'patch-matrix' compartments in the striatum, which provide for separate pathways from the cortex through the striatum to the dopamine neurons in the midbrain and to the output neurons of the basal ganglia. Finally, a description of the transformation that occurs as a result of the organization of striatal outputs into two main output streams, the so-called direct and indirect striatal output pathways to the output neurons of the basal ganglia in the substantia nigra and entopeduncular nucleus (internal segment of the globus pallidus). The function of this organization appears to transform the excitatory inputs from the cortex to the striatum into antagonistic inputs to the output neurons of the basal ganglia. The relative activity in the two striatal output streams thus determines the activity of the output of the basal ganglia. Among the mechanisms that regulate the relative activity in the two striatal output systems is the nigrostriatal dopamine feedback.

9. RELATIONSHIP BETWEEN CORTEX AND BASAL GANGLIA

The major input to the basal ganglia is that from the cerebral cortex to the striatum. This input is multi-faceted in that there are multiple determinants to its organization. First, there is a topographic organization in corticostriatal inputs that is continued 409

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through the circuits of the basal ganglia. Second, the mapping of cortical inputs through the basal ganglia circuitry does not represent a simple point to point map in that there is overlap, or convergence at each level of the system. The nature of this convergence will be discussed. 9.1. TOPOGRAPHIC ORGANIZATION At each level of the basal ganglia there is an apparent topographic organization in the projections from one level to the next. Thus, the cortical projection to the striatum, the projection of the striatum to the globus pallidus, and the projection of the entopeduncular nucleus (internal segment of the globus pallidus in primates) and substantia nigra to the thalamus, each display a topographic organization in which the spatial organization of the source area is maintained in the projection pattern to the target. However, as will be described further, there are other types of connnections that do not adhere to a strict point-to-point topographic mapping, which are overlain on the general topographic mappings that are seen at each level of the system. A variety of studies of the organization of cortical input to the striatum have described the topographic organization of this system in both the rodent (Beckstead 1979; Berendse et al. 1992a; McGeorge and Faull 1989; Webster 1961) and primate (Goldman and Nauta 1977; Kunzle 1975, 1977, 1978; Kunzle and Akert 1977; Selemon and Goldman-Rakic 1985). In this context topography refers to the maintenance of the relationship between cortical areas in the mapping of their projection fields within the striatum. For example, limbic cortical areas, including the hippocampus, piriform and infralimbic cortices, and amygdala, in as much as it might be considered a cortical area, provide inputs to the ventral striatum, including the nucleus accumbens. The prelimbic cortex, dorsal to the infralimbic cortex, provides input to the medial bank of the striatum. This topographic organization is maintained moving from medial to lateral along the cortex, with projections maintaining their medial to lateral relationships in the projections to the striatum. In general limbic, or allo- and peri-allocortical areas project to the ventral striatum, whereas neocortical areas project to the dorsal striatum. Studies of the projections from the striatum to the globus pallidus and to the substantia nigra have shown a distinct topographic organization, in both the rodent (Gerfen 1985; Groenewegen et al. 1993; Groenewegen and Russchen 1984) and primate (Cavada and Goldman 1989b; Cowan and Powell 1966; DeVito and Anderson 1982; DeVito et al. 1980; Flaherty and Graybiel 1993a; Hedreen and DeLong 1991; Nauta and Mehler 1966). Both the striatopallidal and striatonigral projections innervate dual zones in each target structure (Chang et al. 1981; Gerfen 1985; Wilson and Phelan 1982), both of which are topographically organized. The dual zones of innervation correspond to subregions within each nucleus that are delineated by both the pattern of afferent input, and the organization of the dendrites of the target neurons within the nucleus (for example see Figure 13). The topography of each of these projection zones is apparent in coronal sections, and shows a general maintenance of the medial-lateral and dorsoventral relationship of striatal projections in the termination patterns in the target nuclei. In general this topography is maintained in the projections from the striatum to the globus pallidus. In the projections to the substantia nigra, there is a maintenance of the medial-lateral relationships in the terminal fields and an inversion of the dorso-ventral relationships such that more ventral regions of the striatum project to the dorsal parts of the pars reticulata and dorsal regions of the striatum project to the ventral pars reticulata. Projections from the striatum to the entopeduncular nucleus are also topo410

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graphically organized. The indirect pathway projections, which include the globus pallidus and subthalamic nucleus, also display both dual projection systems and topographic organization. It must be stressed that while topographic relationships are apparent in striatal output organization, there are additional features in the organization of these systems. As in the projections of the cortex to the striatum, the topographic organization is not strictly adhered to and there is some overlap of projection fields (Gerfen 1985). Whereas the topographic organization is readily apparent in the medio-lateral and dorso-ventral axes, in the rostro-caudal axis the projections from any given part of the striatum have an extensive rostro-caudal spread. In addition to the dual projection systems of striatopallidal and striatonigral projections, there is a segregation of projections from the 'patch-matrix' compartments. The dual projection systems and striatal 'patch-matrix' compartments will be discussed in detail below. The output neurons of the basal ganglia, which are the GABA neurons in the entopeduncular nucleus and substantia nigra pars reticulata (Mercugliano et al. 1992; Oertel and Mugnaini 1984; Pasik et al. 1988; Ribak et al. 1979), provide inputs that are topographically organized in their projections to the thalamus and superior colliculus in the primate (Fen61on et al. 1990; Ilinsky et al. 1985). In the rat the details of this topography are difficult to work out due to the small size of the structures involved, nonetheless, several studies employing a variety of methods have described a topographic organization (Deniau and Chevalier 1992; Gerfen et al. 1982). Thalamic nuclei innervated by basal ganglia outputs project back to frontal cortical areas in a topographically organized manner in both the primate (Holsapple et al. 1991; Kievet and Kuypers 1977; Schell and Strick 1984) and rodent (Groenewegen 1988). In the rodent these thalamic nuclei include the ventral lateral (VL), paralaminar mediodorsal (MD) and ventral medial (VM) nuclei. The ventral lateral nulceus, which receives input from the entopeduncular nucleus projects back upon the motor cortex. The paralaminar part of the mediodorsal thalamic nucleus projects back upon medial agranular and 'prefrontal' cortical areas. The ventral medial thalamic nucleus projects in a distributed manner to layer 1 of most of the frontal pole (Herkenham 1979). Other thalamic nuclei innervated by basal ganglia outputs, which include the intralaminar nuclei and lateral habenula, provide feedback projections to the striatum (Beckstead 1984; Berendse et al. 1988; Gerfen et al. 1982; Herkenham and Pert 1981). The continuation of the topographic organization of the corticostriatal system through the circuits of the basal ganglia and eventually onto the thalamus has been described in some detail by Alexander, Strick and DeLong from their work in primates (Alexander and Crutcher 1990; Alexander et al. 1986, 1990). They describe 5 parallel corticostriatal systems related to the major cortical areas of origin which are maintained as semi-segregated parallel pathways through the striatum, both segments of the globus pallidus and substantia nigra. These include circuits originating in motor (premotor and supplementary motor areas), occulomotor (frontal eye field), dorsolateral prefrontal, orbital prefrontal and anterior cingulate cortical areas. Each of these cortical areas provides inputs to corresponding regions of the striatum, which for the most part are segregated from one another. Evidence for the maintenance of the topography of corticostriatal projections through the basal ganglia back through the thalamus to the cortex, comes from studies in the primate by Strick and his colleagues (Holsapple et al. 1991; Hoover and Strick 1993; Schell and Strick 1984; Strick 1985). In one study in which they employed anterogradely transported trans-neuronally transferred viruses as axonal connectional markers which 411

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allow for analysis of the projections of striatal neurons which recieve inputs from defined cortical areas. These studies demonstrated that injections of virus into separate frontal motor cortical areas resulted in labeling of virus in distinct zones of the putamen, and secondarily in distinct zones in both the external and internal segments of the globus pallidus. In another study, Hoover and Strick (1993) injected virus which is retrogradely transported and trans-neuronally transferred into separate frontal motor cortical areas, the primary motor cortex, the premotor area and the supplementary cortical area. The 412

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Fig. 20. Diagram illustrating the concept of the maintenance of a general topographic organization (the maintenance of the spatial organization of the projection of one brain area to another) through the connections of the basal ganglia. Projections from the cerebral cortex to the striatum (cortico-striatal) maintain the spatial relationships within the cortex in the targeted regions in the striatum. Similarly the projections of the striatum to the globus pallidus and substantia nigra/entopeduncular nucleus (EP) maintain the spatial relationships of the striatum in the pattern of inputs to these nuclei. Of note are the dual projection zones of the striatum to both the globus pallidus (GP1 and GP2) and substantia nigra (SNrl and SNr2), such that each of these target nuclei contain two topographically organized maps of striatal inputs. Within the substantia nigra the maintenance of the general topography in basal ganglia connections is evident in the projection of the entopeduncular nucleus (EP) to the ventral lateral thalamus (vl) and of the substantia nigra projection to the ventral medial (vm) and mediodorsal (md) thalamic nuclei. It is important to stress that the topography depicted is only a general organizational feature, the borders in the projections are not precisely delimited, and there are many examples of connections that do not correspond to the topographic organization at all (i.e., the widespread projections in the corticostriatal projections, particularly in the rostral-caudal axis). (

arm representation region of each cortical area was injected. In each case virus was identified in the globus pallidus, having been transported retrogradely to neurons in the thalamic nuclei which project to these cortical areas, the ventrolateral oralis and area X, trans-neuronally transferred and transported in the projection axons of these neurons to the internal segment of the globus pallidus. Trans-neuronally transported virus was localized in spatially separate regions of the internal segment of the globus pallidus. The region in which virus was identified in the globus pallidus from each cortical area was distinct, and topographically related to the cortical area of origin. Together these studies suggest a segregation and rather strict maintenance of the topographic organization of parallel organization of cortical outputs, which is maintained through the basal ganglia circuits to thalamic feedback to the cortex. The topographic organization of the cortical projection to the striatum and the continuation of this organization through each successive level of the basal ganglia circuitry is diagrammed in Figure 20. Although this diagram represents the organization in the rat, a similar organization applies to the primate. It must be emphasized that the regional boundaries indicating target zones of cortical inputs are not absolute and that there is some overlap of projection fields. The organization of the overlap of projection fields will be discussed in some detail below. 9.2. O V E R L A P O F INPUTS: C O R T I C O - C O R T I C A L O R G A N I Z A T I O N The topographic organization of the cortico-striatal system, which is carried through the striato-pallidal, striato-nigral, pallido-thalamic and nigro-thalamic pathways, is fairly well established. This principle of basal ganglia organization is the basis of the concept of parallel pathway loops connecting functional cortical regions through the basal ganglia with thalamic nuclei that project back upon frontal cortical areas (Alexander et al. 1986). However, this organization does not reflect a point to point mapping of cortical areas with the basal ganglia circuit. Another principal organizational feature of the corticostriatal system is the fact that a given cortical area projects to a domain of the striatum that is proportionately larger than the cortical area of origin, and so implies considerable divergence and convergence in the corticostriatal projection. This divergence is particularly extensive in the rostro-caudal dimension (Selemon and G o l d m a n - R a k i c 1985; Yeterian and Hoesen 1978). Yeterian and Van Hoesen (1978) made the observation that the parietal cortex and prefrontal cortex provide inputs that appear to overlap over a fairly extensive rostro-caudal area. As these areas are con413

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C.R. Gerfen and C.J. Wilson

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Fig. 21. Diagram to illustrate a reconciliation of two concepts of basal ganglia organization related to the corticostriatal system, the existence of parallel closed loop circuits from cortical areas through the basal ganglia back to the cortex (Strick et al., 1985) and the widespread and often discontinuous projections of the cortex to the striatum (Kunzle, 1975; Yeterian and van Hoesen, 1978; Flaherty and Graybiel, 1993). Depicted are two cortical areas (area 1 and 2) each which is organized in a somatotopic manner (a,b,c,d and A,B,C,D). The two cortical areas are interconnected with connections between homologous representations in each cortical area. Each cortical area provides an input to a topographically related zone in the putamen, cortical area 1 to a dorsal area and cortical area 2 to a more ventral area. The spatial relationship between the two cortical areas is maintained in the projections to the external (GPe) and internal (GPi) segments of the globus pallidus and in the projections of the GPi to the thalamus and back to the cortex. The maintenance of the spatial projection fields from cortex to striatum to globus pallidus to thalamus and back to cortex forms the basis of the parallel closed loop model. Each cortical area does not provide inputs to the entirety of the recipient area in the putamen, rather it provides input to the subfield of the putamen that corresponds to the topographically related part of the cortical area of origin. In addition, each cortical area provides a secondary input to the homologous region of the topographically related field of the other cortical area. Thus, subarea 'A' of cortical area 1 provides inputs to subarea 'A' of putamen area 1 and subarea 'a' ofputamen area 2. Thus, each cortical area provides a striatal input that has the appearance of being discontinuous as they map onto the appropriate subareas of each target area in the putamen. Such organization is continued through the rest of the basal ganglia circuits. nected by cortico-cortical connections they suggested that cortico-striatal organization is r e l a t e d t o t h e c o r t i c a l c o n n e c t i o n s o f t h e a r e a f r o m w h i c h t h e c o r t i c o - s t r i a t a l i n p u t s arise. T h e y f o r m u l a t e d a r u l e w h i c h s u g g e s t e d t h a t a r e a s o f c o r t e x w h i c h a r e i n t e r c o n n e c t e d b y c o r t i c o - c o r t i c a l c o n n e c t i o n s p r o v i d e o v e r l a p p i n g i n p u t s to t h e s t r i a t u m ,

414

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whereas areas of cortex that are not interconnected do not. Theirs was the first formulation of a concept to explain the widespread nature of cortico-striatal projections from a given region. The rule suggested by Yeterian and van Hoesen (1978) has been examined in subsequent studies with mixed results. Selemon and Goldman-Rakic (1985) confirmed that both parietal and prefrontal cortical areas provide inputs to the striatum that extend over a large expanse in the longitudinal axis, but they focussed on areas of interdigitation and not the areas of overlap in the cortico-striatal projections from these interconnected cortical areas. They concluded that areas of cortico-striatal convergence are not related to patterns of cortical connectivity. However, subsequent, more detailed mapping of the interconnections between parietal and prefrontal cortex (Andersen 1990; Cavada and Goldman 1989a; Cavada and Goldman 1989b) called for a reassessment of this conclusion (Cavada and Goldman 1991). Connections between posterior parietal cortex and prefrontal cortex were shown to have a precise pattern of segregated connectivity, which reflects modality or functionally specific connections. Thus, the posterior parietal area 7a and lateral intraparietal area (LIP), which have visual and visual-motor functions, are interconnected selectively with prefrontal areas 46 and 8a, respectively, whereas, parietal area 7b, which has somatosensory functions, is connected with prefrontal area 45 (Andersen et al. 1990). The specificity of these associational connections was not taken into account in the Selemon and Goldman-Rakic (1985) study. For example, their injection of tracer into the posterior parietal cortex included an extensive area that projected to a larger domain of the prefrontal areas than was injected with the second tracer. Consequently, if the simple concept that cortically connected areas provide overlapping striatal inputs were correct, the result they obtained, with areas of overlap and areas of non-overlap, would be expected. Cavada and Goldman-Rakic (1991) did in fact analyze cortico-striatal inputs in this context and came to the conclusion that the regional distribution of parietal and prefrontal corticostriatal projections did in fact reflect the interconnections between functionally related cortical areas. The concept that functionally interconnected parietal and prefrontal corticostriatal inputs are directed to overlapping regions of the striatum (Yeterian and van Hoesen 1978; Cavada and Goldman-Rakic 1991) suggests that the extensive longitudinal distribution of cortico-striatal inputs is related to cortico-cortical connections. However, in these studies the detailed patterns of overlap and interdigitation that are observed in experiments in which the corticostriatal projections from different cortical areas are examined with separate tracer injections into these areas in the same animal (Selemon and Goldman-Rakic 1985), leaves open the question of the relationship between corticocortical and cortico-striatal organization. The problems of relating the organization of cortico-cortical connections to corticostriatal organization is confounded by the complex connection patterns in each of these systems. Although it is true, as described by Yeterian and Van Hoesen (1978), and later by Selemon and Goldman-Rakic (1985) that cortical areas provide inputs that extend over a considerable rostro-caudal domain, the innervation patterns are by no means uniform. In many instances projections from a given cortical area show distributed but discontinuous patterns of input to the striatum. Examples of such discontinuities have recurred in the literature beginning with studies of Kunzle in the 1970's (Kunzle 1975, 1977). Particularly striking are the multiple representation zones within the striatum from somatosensory and motor cortical areas. Based on functional mapping studies employing 2-deoxyglucose Brown has suggested that multiple innervation patterns in the striatum from somatosensory cortical areas reflect multiple somatotopically organ415

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ized convergence zones in which cortical inputs from different functional modalities, such as motor and somatosensory areas, converge in a combinatorial manner (Brown 1992; Brown and Feldman 1993). The discovery of discontinuous zones of innervation in the projections from somatosensory and motor cortex (Kunzle 1975, 1977) made it possible to exploit the internal organization of these areas to design more detailed studies of the corticostriatal system. These studies have made use of the precise body map contained in motor and somatosensory areas to study the corresponding parts of two different cortical areas, something that could not be accomplished in studies of posterior parietal and prefrontal cortical regions. Graybiel and her colleagues have examined the connectional basis of this organization in a set of careful studies in which they examined the relationship of cortico-striatal inputs from cortical areas that had been mapped in terms of somatosensory and motor function (Flaherty and Graybiel 1991; Flaherty and Graybiel 1993b; Parthasarathy et al. 1992). In one study the projections from different somatosensory areas were examined (Flaherty and Graybiel 1991; Flaherty and Graybiel 1993b). Electrophysiologically mapped regions of areas somatosensory areas 3a, 3b and 1 were injected with anterograde tracers (Flaherty and Graybiel 1991). They found that injections into matched body part represenation sites in different somatosensory areas provided inputs that overlapped in their projections into the striatum, and that these projections displayed multiple innervation zones. Conversely, injections into different body part regions of cortical area S1 provided inputs to multiple non-overlapping striatal regions. In another study Parthasarathy et al (1993) examined the cortico-striatal projections of two frontal cortical areas that are involved in eye movements, the supplementary eye fields and the frontal eye fields. They found that the degree of overlap of corticostriatal inputs from injections of tracer into these cortical areas was directly correlated with the degree of cortical connectivity between the injected areas. Similar to the organization of somatosensory cortical inputs, there were multiple innervation zones within the striatum from these motor cortical areas. When striatal inputs from non-occulomotor supplementary motor cortex were compared with those from frontal eye fields, there was neither an overlap of inputs in the striatum nor was there evidence of interconnections between the cortical areas injected. In a third study the organization of corticostriatal projections of somatosensory (S 1) and primary motor (M 1) projections to the striatum were investigated in the squirrel monkey (Flaherty and Graybiel, 1993). They found, as had been reported before, that injections of tracers into each of these regions provided inputs that are directed to the putamen and distributed in multiple, discontinuous zones. What they also found was that when somatotopically related areas of M1 and S1, such as the hand representation, that the discontinuous zones of each cortical projection zone overlapped in the ipsilateral putamen. This result suggests that each of these cortical areas provide multiple somatotopically organized cortico-striatal projections and that within the striatum, the multiple somatotopically organized regions receive convergent input from each cortical area. However, they also reported that the contralateral projections did not display this same pattern of overlap between homotypic somatotopic zones of the cortex. This led them to suggest that 'neither patterns of cortical connectivity nor homotypical relationships are infallible predictors of corticostriatal overlap'. Graybiel and her colleagues have provided some thoughtful discussion as to the functional implications of their results (Flaherty and Graybiel 1991, 1993b; Parthasarathy et al. 1992). They suggest 'that whether inputs from particular cortical regions converge in the striatum depends on aspects of their functions, which are only sometimes 416

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mirrored by their cortical connectivity.' They suggest further 'that functionally related inputs would converge, with the degree of functional relatedness being determined by the striatal target. This would allow for activity dependent sculpting of the convergence patterns. For example, if the signal sent to the superior colliculus via a striato-nigraltectal connection coded for saccades without respect to body movements, it would be reasonable to have converging inputs from cortical eye field zones to the striatal origin of the pathway, but not ... from ... areas encoding nonoculomotor body movements. By contrast, targeting to output channels consolidating somesthetic signals might call for convergence of cortical inptus, for example deep and cutaneous inputs form areas 3a and 3b, respectively, whether the cortical areas are connected or not.' They suggest that the existence of functionally organized zones within the striatum which, because these zones are discontinuous areas within the striatal matrix, they term 'matrisomes'. According their ideas, the determinant of the overlap of cortical inputs is related to the function of the output of the 'matrisomes'. Thus, they suggest that the functions of 'matrisomes' represent combinations of cortico-striatal inputs that are different from the combinations of cortico-cortical inputs to cortical areas. This latter conclusion has important implications in the context of the functional organization of the striatum, and in particular with the concept of parallel functional cortical-basal ganglia circuits as put forth by Alexander and his colleagues (Alexander and Crutcher 1990; Alexander et al. 1986, 1990). The basis of the 'matrisome' concept is that the mulitple innervation zones provided from a single cortical site, represent functional units in which cortical information is reorganized. Such discontinuous patterns of corticostriatal inputs are clearly established from a number of studies employing a variety of tracers in a number of species over an extended period of time. However, the basis of such patterns is still open to study. The existence of multiple discontinuous zones of corticostriatal innervation, and the combinational convergence of inputs within them, cannot be understood at the tissue level. Some knowledge of the innervation patterns of single corticostriatal cells is required. If every corticostriatal axon arising from a small region of cortex projects to every one of these multiple discontinuous zones, then they represent a single topographic representation that is strangely shaped and nothing more. Each of the zones from one cortical area will have basically the same pattern of convergence as every other. If, on the other hand, individual corticostriatal neurons innervate single matrisomes, then they represent parallel independent output pathways from the cortex. Each set of corticostriatal neurons, specialized for projecting to a single matrisome, could carry slightly different information destined for convergence with information from a different area. The cellular heterogeneity of the corticostriatal projection could find useful work in this scheme, with different corticostriatal cell types from a single cortical region projecting to different matrisomes and converging with a specific cell type from a different cortical region. Even if there were not specific cell types projecting to each of the matrisomes innervated by a cortical region, each matrisome could have a unique functional identity from the point of view of the cortical regions innervating it. Between these extremes are a variety of intermediate schemes. For example, one matrisome (or one set of matrisomes) might be innervated by all corticostriatal neurons in a region. This could be considered the primary striatal recipient zone of that cortical region. Other matrisomes would then be innervated by collaterals from a subset of the axons, forming a secondary set of recipient zones, each with a different subset of corticostriatal innervation. Convergence of inputs from cortical regions could be organized so that each cortical region had a primary recipient zone in the neostriatum, and secondary zones that overlap the 417

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primary zone of a set of other cortical areas. A scheme of this sort could reconcile the data for parallel independent pathways through striatum and that suggesting convergence of cortical inputs. Cellular heterogeneity of the corticostriatal projection could also help to explain the absence of a single simple rule governing the convergence of corticostriatal inputs. Rules based on cortico-cortical connections, like that suggested by Yeterian and van Hoesen (1978) already raise the issue of cellular heterogeneity, because there are a variety of different kinds of cortical neurons that form corticocortical connections. Some of these also make corticostriatal connections. If, for example, only those corticocortical neurons which participate in forward projections (that is, those primarily directed at layers 3 and 4) participate in making overlapping corticostriatal fields, one would expect to see overlapping corticostriatal projections of some interconnected cortical fields but not others. Likewise for the backwards and lateral projections. A variety of combinations are possible given the variety known to exist in corticocortical projections. The history of study of the basal ganglia consists largely of conjecture about different functional modalities combining in a unique manner in the basal ganglia. For example, Nauta suggested that the integration between the limbic and motor systems occured as a consequence of the projections from the ventral 'limbic' striatum, to the dopamine neurons which provide feedback to the dorsal 'non-limbic' striatum (Nauta et al. 1978). One of us suggested that the patch-matrix organization accomplished this integration (Gerfen 1984). However, upon more detailed analysis it now appears that input to the patch compartment is not unique to the limbic cortex, but is a component of non-limbic cortices as well (Gerfen 1989). As will be discussed later, the most recent data suggests that the patches (striosomes) in the striatal regions that receive inputs from non-limbic cortical areas do not receive inputs from limbic cortical areas, but instead are innervated by corticostriatal neuron types that are present in both kinds of cortical areas but are most numerous in the limbic cortex (Gerfen 1989). The emphasis on convergence of information in the striatum has meanwhile shifted to the discontinuous zones of projection in the matrix (matrisomes). The question of organized convergence of information in the matrix continues to be discussed along the same well-established lines. It should be considered that despite the predisposition to explain the basal ganglia as a mixing place for inputs, the unique contribution of the basal ganglia may not arise from the formation of combinations of cortical inputs, but rather from the nature of the computational operation performed on these inputs by striatal neurons. Some of its operations may be performed implicitly by the connectivity of its output system. Later we will describe two features of that connectivity, one related to the separation of the 'direct' and 'indirect' pathways from the striatum to the output nuclei of the basal ganglia, and the other being the 'patch-matrix' compartments. But it should not be overlooked that the intrinsic connections of the striatum, and the electrical properties of the striatum equip it to perform operations on its input that could not be reproduced by the cortical circuitry, even if it were receiving exactly the same afferent information. This is, of course, the essence of the parallel circuit model put forward by Alexander and his collegues (Alexander and Crutcher 1990; Alexander et al. 1986, 1990). 9.3. STRIATAL OUTPUT SYSTEMS: TOPOGRAPHY / CONVERGENCE / DIVERGENCE The topographic organization of cortical inputs to the striatum, of striatal outputs to the globus pallidus (external segment), internal segment of the globus pallidus (en418

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topeduncular nucleus), and substantia nigra, and from these output nuclei of the basal ganglia to the thalamus is the basis of the multiple segregated loop circuit model of cortical-basal ganglia organization (Alexander et al. 1986). On the other hand, as discussed above, there is clear evidence of convergence of cortical inputs to the striatum. Studies of the organization of the outputs of the striatum reveal a similar organization, namely that there is a clear topographic organization plus patterns of convergence (Gerfen 1985). A question that is raised is how these two schemes, topography and convergence, might be reconciled. Hoover and Strick (1993) have shown, using retrogradely trans-neuronally transported virus tracing, that the projections from the output nuclei of the basal ganglia (the internal segment of the globus pallidus) are organized such that the topographic organization within the globus pallidus is maintained through the thalamus back to frontal cortical areas. This suggests that regardless of the organization of the cortical and striatal projection systems that the output organization of the basal ganglia reflects the organization of the frontal cortical areas. Strick and his colleagues argue that the projection from the cortex through the basal ganglia also reflect this organization in that the cortical inputs likewise maintain their topographic organization. Overlain on this topographically organized backbone of cortico-basal ganglia circuitry are convergent (and/or divergent) components. Thus, as demonstrated by Graybiel and her colleagues, somatotopically similar parts of different cortical areas provide inputs to overlapping striatal regions (Flaherty and Graybiel 1993b), which in turn reconverge in the projections of the striatum to the segments of the globus pallidus (Flaherty and Graybiel 1993a). This might be considered to represent a reorganization of cortical information in the patterns of divergence of cortico-striatal inputs and reconvergence in the striato-pallidal projections. However, a more straightforward model would suggest that these patterns represent a topographically organized system which is overlain with secondary connections that represent a mapping in the striatum of cortico-cortical connectivity. This organization is schematically diagrammed in Figure 21. The studies of Strick and Graybiel have focussed on the organization of the connections of cortical areas that provide their major inputs to the putamen. These cortical areas are for the most part those associated with primary motor and somatosensory cortical areas or those cortical areas with inputs to these areas, including premotor and supplementary cortical areas. One of the reasons for studying such areas is that they are organized in a somatotopic manner, which aids in a precise mapping of the projections relative to physiologically defined determinants. Other cortical areas, whose functions are somewhat removed from a direct relationship to defined movement or sensory functions, such as prefrontal or parietal cortical areas, have more widespread patterns of cortico-cortical connectivity. These areas, generally termed associational cortical areas, provide less precisely organized inputs to the striatum, or less precise in patterns that we now recognize. Most likely these inputs might be described as less precise because the functional organization of the cortical areas have not been as well characterized. The corticostriatal inputs from these areas are marked by extensive distribution patterns that display a topographic organization in the medio-lateral axes, but also display extensive distributions in the rostro-caudal axis. Similarly, the projection of the striatal regions that receive these inputs display the same pattern of organization in their projections to the globus pallidus and substantia nigra, a maintenance of the mediolateral topography, with extensive distribution in the rostro-caudal axis. This organization has been observed in both the primate and rodent. Examples of the projections from the striatum to the substantia nigra in the rat are shown in Figure 22 to illustrate the 420

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extent of the rostro-caudal spread of the striatal projection system. Again, this suggests that there is a similar organization in the projections from the cortex to the striatum and in the projections of the striatum. Within these patterns of projection another aspect of the organization of striatal projections is apparent, that is the dual target zones within the substantia nigra. This will be discussed below. The topographic organization of cortical and basal ganglia circuitry has been well documented in both rats and primates. However, as has been discussed for corticostriatal inputs, the organization does not represent a point to point mapping at each stage of the system. The organization of cortico-striatal inputs provides some convergence of inputs from different cortical areas. The possible convergence at other levels of the basal ganglia has been proposed. For example, it has been suggested that the spread of the dendrites of neurons in the globus pallidus and substantia nigra are so extensive that they provide for a convergence of inputs from widespread regions of the striatum. According to this view, the topographic organization of striatal outputs is superseded by the convergence that is a consequence of the dendrites of the target neurons. While we would not argue for a strict topographic organization, there are several reasons to suggest that the extent of convergence of inputs as a consequence of the dendritic spread must be considered with some caution. As has been described the input of individual striatopallidal axons make multiple contacts with individual pallidal neuron dendrites, which appear to ensheath these target dendrites. This feature of synaptic organization is contrasted with that of cortical inputs to the striatum, where individual corticostriatal axons may make only single contacts with many spiny neurons. This organization typifies inputs that might be considered to be transferring input from single cortical neurons to multiple striatal neurons. On the other hand, in the globus pallidus, there appears to be an organization that suggests a rather tighter transferance between striatal neurons and a limited number of pallidal target neurons. There are two other distinctive features of striatal output organization. The first is the segregation of the projections of the 'patch-matrix' compartments of the striatum. This will be discussed in a later section. The second, which has been mentioned, is the dual projection zones of striatal inputs in both the globus pallidus and substantia nigra. As will be described these dual projection systems appear also in the organization of other components of the basal ganglia, including the projection of the subthalamic nucleus. 9.4. STRIATAL OUTPUTS IN RELATION TO NIGRAL OUTPUTS: DUAL OUTPUT SYSTEMS A distinctive feature of striatal output organization is the dual projections from the striatum to subdivisions of the globus pallidus and substantia nigra (Chang et al. 1981; Gerfen 1985; Wilson and Phelan 1982). This organization has also been observed in the primate (Parent and Hazrati 1993). Striatal projections to the globus pallidus have extensive axon arborizations in a region immediately adjacent to the striatum, and a second arborization zone in the central part of the globus pallidus. In the case of the striatopallidal projection, the dual projections have been demonstrated to arise from individual neurons (Chang et al. 1981). The dual striatonigral projection targets a region in the dorsal region of the substantia nigra pars reticulata, and a second zone that lies immediately above the cerebral peduncle. It has not been demonstrated that individual striatal neurons contribute projections to both zones of the pars reticulata, although this is likely. At the least they arise from within the striatal matrix and from very closely 421

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associated neurons. These dual projection systems are not to be confused with the patch-matrix projections. The dual nature of inputs to the globus pallidus and substantia nigra is not only observed in the striatal projections to these nuclei. Kita and Kitai (1987) have also observed a similar organization in the projection of the subthalamic nucleus to these nuclei. The projection patterns charted in their study bear a remarkable resemblance to those from the striatum. This suggests that this aspect of the organization of basal ganglia circuits is maintained not only in the organization of striatal outputs but also in the organization amongst the nuclei that are the targets of this striatal projection. In both the globus pallidus and in the substantia nigra the dendritic morphology of neurons in these nuclei conform to the dual innervation patterns from the striatum 422

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Fig. 23. Illustration of the dual projections from A) the striatum (CP) to the globus pallidus (GP) and substantia nigra pars reticulata (SNr) and from B) the subthalamic nucleus to the globus pallidus (GP) and substantia nigra pars reticulata (SNr). In each system afferents target the same two regions in the GP, an area immediately adjacent to the striatum and second area more medial, and the same two regions in the SNr, an area medial and dorsal adjacent to the substantia nigra pars compacta and a second area situated ventrally against the cerebral peduncle. The dual target zones in both the GP and SNr have neurons whose dendrites appear to conform to the pattern of afferents to these regions (see Fig. 13 and 18). In addition individual striatal neurons and individual subthalamic neurons provide collaterals to both regions in each nucleus. C. Sagittal diagram of projections from substantia nigra to the superior colliculus. Neurons in the two subregions of the substantia nigra pars reticulata (SNr) that are defined by the pattern of striatal and subthalamic afferent inputs project to different parts of the superior colliculus. Dorsal SNr region neurons (white) provide input to the rostral superior colliculus. Ventral-caudal SNr region neurons (black) provide input to the caudal superior colliculus. D. A top view diagram of the superior colliculus on which is depicted the organization of the eye movement saccades that are generated by stimulation of the intermediate layer. Longer saccades are generated in the caudal superior colliculus, shorter saccades are generated in the rostral superior colliculus, and the most lateral rostral zone is involved in fixation. The organization of afferents from the dorsal SNr (white), directed to the rostral, short saccade and fixation region of the superior colliculus, and from the ventral caudal SNr (black), directed to the longer saccade region of the superior colliculus are depicted.

(Gerfen 1985). Thus in the globus pallidus, neurons in the region that is immediately subjacent to the striatum have dendrites that are distributed in a pattern that conforms to a 'shell'-like region of the globus pallidus, whereas neurons in the central region of the globus pallidus are distributed in the central region and do not appear to extend into the pallidal 'shell' region (Kita and Kitai 1994). Similarly, in the substantia nigra there are two zones of neurons in the pars reticulata, ignoring the d o p a m i n e neurons in the pars reticulata. Again, as in the globus pallidus there is one region that forms a 'shell' like structure, in this case forming a region immediately above the cerebral peduncle, and a dorsal zone region that is the region between the ventral 'shell' region and the pars compacta. N e u r o n s in these two regions have dendrites that are distributed so as to conform with the shape of the regions ( G r o f o v a et al. 1982). This organization was first described by G r o f o v a et al. (1982) based on the m o r p h o l o g y of the dendrites of pars reticulata neurons. The organization of the substantia nigra pars reticulata into subregions appears not only to be related to the organization of inputs from the striatum and subthalamic nucleus, but in the organization of its outputs. The organization of the projections of the substantia nigra pars reticulata to the thalamus and to the superior colliculus appear to maintain a rough topography. This topographic organization has been described by Gerfen et al. (1982) and in considerable detail by Deniau and Chevalier (1991). Thus, projections to the ventral medial, mediodorsal, and intralaminar thalamus, as well as those to the projections to the superior colliculus, display a topographic organization. This t o p o g r a p h y involves both the central and peri-peduncular 'shell' region of the pars reticulata neurons. N e u r o n s projecting to a particular topographically related part of any of these structures are organized in one of the two pars reticulata regions. This organization of the nigral o u t p u t neurons was described by Deniau and Chevalier (1991) to have the appearance of distinct lamellae, much like that of an onion. This organization has been remarked upon repeatedly, by Gerfen et al. (1982), Deniau and Chevalier (1991), and Redgrave et al. (1992). The functional signficance of the dual projection systems of striatal outputs may be related to the organization of the target structure of the nigral outputs. In terms of the organization of the substantia nigra projection to the superior colliculus it appears that neurons in the dorsal region project to the rostral superior colliculus, whereas those in 423

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the peripeduncular 'shell' project to more caudal regions. In addition, each of these nigro-tectal projections maintains a medio-lateral topography. Redgrave et al. suggested that the organization of the nigro-tectal organization reflects differences in the of both afferent and efferent organization of the intermediate layer of the superior colliculus that is the target of these inputs (Redgrave et al. 1992). An alternative organization within the superior colliculus that might be the basis for the organization of the nigro-collicular pathway may be related to the map of eye and head movement generation. Neurons in the intermediate layer of the superior colliculus appear to be involved with the generation of eye and head movements. Within this layer, movements generated by stimulation are mapped in an orderly manner such that small saccades are produced by stimulation in the rostral half of the colliculus and larger saccades, accompanied by head movements are mapped in the caudal half. At the rostral-lateral pole of the superior colliculus is a zone which is involved in fixation (Munoz and Wurtz 1992, 1993). This map within the superior colliculus conforms, at least roughly, to the organization of the outputs from the two zones within the substantia nigra pars reticulata. Whether there exists a similar organization of dual outputs from the substantia nigra and from the internal segment of the globus pallidus to the thalamus remains to be determined. In this context the results reported by Hoover and Strick (1993) are compelling. In their study in which virus injected into the cortex was retrogradely and transneuronally transported to the internal segment of the globus pallidus in primates they reported that in addition to the topographic organization of the virus labeling, there were two zones within the nucleus, that correspond to the dual innervation zones from the striatum. This would suggest the existence of dual outputs from the internal segment of the globus pallidus to the thalamus, which in turn converge on particular cortical areas. The organization of the dual pallidal input to the thalamus and the organization of dual thalamic projections back to the cortex remains to be worked out. Several possibilities might be investigated. One is that the dual pallidal outputs to the thalamus might innervate different compartments within the same thalamic target nucleus, in this case VLo and area X. Studies of the projections of the pallidum to this nucleus have revealed a non-homogeneous innervation pattern (Holsapple and Strick, 1991), which is similar in organization to that described by Jones and co-workers for the organization of other ventral thalamic relay nuclei (Rausell et al. 1992; Rausell and Jones 1991a,b). It is possible that one part of the internal segment of the globus pallidus innervates one of the compartments, whereas the other zone innervates the other compartment. If VLo is organized in a similar manner to VPL and VPM, then as described by Jones, these different compartments might project back to different layers of the same cortical areas. Alternatively, the dual pallido-thalamic output might target different thalamic nuclei, VLo and the centro-median thalamic nucleus. In this case these two different thalamic nuclei also provide convergent inputs to the same cortical areas, but again to different laminae. While further work needs to be done to establish the specific functional organizational significance of the dual projection zones in the striatal output systems there are several determinants of this system that are now clear, and distinguish this organization from other aspects of striatal output organization. First, individual striatal neurons innervate both zones of the external segment of the globus pallidus. It is also likely that individual striatal neurons also innervate the two zones of the internal segment of the globus pallidus and the substantia nigra as well. This feature of the dual projection systems is distinct from that of the output organization of the patch-matrix striatal compartments, 424

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which might also be viewed as providing dual projection systems to separate regions of the target nuclei. As will be discussed below, the dual projections from the patch-matrix compartments arise from separate neuron populuations. Second, the separate zones of the substantia nigra provide topographically organized inputs to the superior colliculus, and a similar organization of nigro-thalamic and pallido-thalamic projections seems likely. However, not only do individual neurons provide input to both zones, but each region of the striatum provides inputs to both zones, such that there are dual topographically organized inputs from the striatum to these target nuclei. Redgrave had suggested that different regions of the striatum might innervate the separate output pathways of the nigro-tectal pathway. However, it would appear that each striatal region provides inputs to both. Third, these dual projection systems are a very prominent feature of basal ganglia organization in rats and primates, suggesting that there is a significant functional purpose for this organization. 9.5. SUMMARY OF ORGANIZATION OF CORTICO-BASAL GANGLIA CIRCUITS As discussed above, there appears to be a general topographic organization in the projections of the cereberal cortex to the striatum, and in the organization of the basal ganglia circuits such that a number of functionally defined cortico-basal ganglia 'loops' may be defined. However, the organization of the basal ganglia does not reflect a precise point to point remapping of the cortical inputs. In particular there is substantial evidence that there exists a convergence of inputs from multiple cortical areas in the striatum such that cortical areas that are interconnected provide convergent inputs to the striatum. The patterns of these convergent inputs remains open to study. Several possibilities have been proposed. On the one hand, Graybiel and her colleagues have suggested that the pattern of convergent inputs to the striatum represents a remapping of cortical systems such that functionally related information from different cortical areas converge on 'matrisomes' within the striatum, which determine the output organization of the striatum. On the other hand, Strick, Alexander and DeLong, have proposed that the organization of functional systems within the cortex are carried through topographically segregated parallel loops through the basal ganglia circuits to feedback onto the frontal cortical areas, from which these loops arise. Strick and his colleagues have provided evidence to support the idea that the output structure of the basal ganglia circuits are organized into segregated loops that feed back through the thalamus in a topographically organized manner. Reconciliation of these two models would be provided by the possible convergence of secondary sites of terminations of corticostriatal inputs, related in part to the organization of cortico-cortical connections, onto primary topographically related cortico-striatal projections. In this reconciled model, there would exist both the primary segregated loop model and convergence of functionally related information from different cortical areas onto these cortical-basal ganglia loops. The organization of the output systems of the striatum, and of their targets appear to reflect that of the cortical inputs. There is a general maintenance of a topographic organization, which is overlain with a patterns of convergence. As in the corticostriatal system, the patterns of convergence are most extensive in the rostrocaudal dimension. In addition, there is an additional organization in the basal ganglia loops, that of dual projection systems, which occur at each level of the projections from the striatum to the globus pallidus and substantia nigra as well as in the intermediate system which includes the subthalamic nucleus. This dual projection system arises from individual neurons at 425

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each stage of the basal ganglia circuit. The functional significance of this dual projection system may be related to the organization of the final targets of the basal ganglia output, the superior colliculus and thalamus.

10. STRIATAL PATCH/MATRIX COMPARTMENTS Within the complexity of the striatum it is important to identify those aspects of organization that provide underlying mechanisms which might account for the heterogeneity of functional structure. For example, the underlying organization of striatal output neurons displays considerable homogeneity in that two major subpopulations may be defined connectionally, by their respective projections to the globus pallidus and to the entopeduncular nucleus/substantia nigra, and neurochemically, by their selective expression of dopamine receptor subtypes and certain neuropeptides (Gerfen 1992). The rather uniform distribution of these neurons in all regions of the striatum underscores the homogeneity of this aspect of striatal organization. However, as will be detailed below, although in some respects the regulation of these two subpopulations also reflects a mechanistic uniformity, there are other aspects of regulation that reveal both regional and subregional heterogeneity. Such heterogeneity is related to compartmentally organized systems that are overlain on the organization of striatopallidal and striatonigral output neurons and function to regulate these output neurons. One such system is the so-called 'patch-matrix' striatal compartments which are involved in the way that the dopamine input to the striatum is regulated (Gerfen 1992). Patch-matrix striatal compartments are often described on the basis of neurochemical markers (Gerfen et al. 1985; Graybiel and Ragsdale 1978; Herkenham and Pert 1981). However, as will be described, in some cases such compartmental heterogeneity reflects regulatory processes, particularly in the relative levels of different neuropeptides (Gerfen et al. 1991). Patch-matrix compartments may be defined precisely on the basis of connections of the neurons in these compartments (Gerfen 1984, 1985, 1989; Gerfen et al. 1987). Such a definition is important to understanding the functional organization of the striatum as it is critical to distinguish the underlying mechanisms that give rise to the different regulatory mechanisms that give rise to heterogeneity within the striaturn. Thus, although we will begin with the organization of the nigrostriatal dopamine system to the patch and matrix compartments, it is important to be forewarned that the underlying organization of these compartments are related to the segregation of cortical inputs that target different populations of striatal output neurons that themselves target different neurons in the other components of the basal ganglia. While some neurochemical markers show regulation-dependent distribution patterns relative to the patch-matrix compartments, most notably the neuropeptides in striatal medium spiny neurons (Gerfen et al. 1991), other neurochemical markers show patterns consistent with the connectional determinants of patch-matrix organization. The first of these to be identified as a patch-matrix marker is the binding pattern to mu opiate receptors, which is greatly enriched in the patch compartment (Herkenham and Pert 1981; Pert et al. 1976). Another is the distribution of axon collaterals of striatal somatostatin-containing interneurons (Gerfen 1984; Gerfen et al. 1985). Particularly useful is the localization of the calcium binding protein calbindin in striatal matrix projection neurons (Gerfen et al. 1985). This marker has been particularly useful as displays the same patch-matrix organization in the striatum in rats and primates (Gerfen et al. 1985). These markers, all show consistent patch-matrix distributions relative to one another 426

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in most regions of the striatum and have been useful in establishing the connectional basis of patch-matrix organization. The distribution of these markers is shown in Figure 24. 10.1. NIGROSTRIATAL DOPAMINE SYSTEM Dopamine innervation of the striatum (Bj6rklund and Lindvall 1984) is relatively dense and when considered in total is rather uniform. However, this belies an underlying organization of the nigrostriatal system into patch- and matrix-directed systems (Gerfen et al. 1987; Gerfen et al. 1987; Jimenez and Graybiel 1987; Langer and Graybiel 1989). The first indication of the compartmental organization of the nigrostriatal dopamine system came from developmental studies which revealed that in the early postnatal striatum dopamine input is distributed in patches, and that during subsequent development innervation of the matrix is completed (Olson et al. 1972; Tennyson et al. 1972). Neuroanatomical tracing studies demonstrated that this developmental sequence is a consequence of the dopamine projection to the patch and matrix compartments arise from distinct sets of dopamine neurons in the substantia nigra (Gerfen et al. 1987). The distribution of dopamine neurons in the ventral midbrain, labeled with tyrosine hydroxylase immunoreactivity, and those that project to the striatum are shown in Figure 25. The midbrain areas in which dopamine neurons are distributed include the ventral tegmental area, which is the ventral medial most region of the midbrain, the substantia nigra, which includes the pars compacta, in which dopamine neurons are

r .

i Fig. 24. Patch and matrix striatal compartments are labeled with neurochemical markers. A) The patch compartment is labeled with 3H-naloxone binding to mu opiate receptors (white in the darkfield photomicrograph). B) The matrix compartment is labeled with calbindin-immunoreactivity, which labels spiny projection neurons that provide inputs to the substantia nigra pars reticulata. The correspondence between calbindinpoor zones (black arrows) and mu opiate binding sites (white arrows) is seen to occur in all regions of the striatum. Calbindin-immunoreactivity is relatively weak in the dorso-lateral striatum, which nonetheless contains opiate receptor patches.

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Fig. 25. The organization of the nigrostriatal dopamine (DA) pathway from the midbrain to the striatum (sagittal diagram at upper right) is diagrammed to show the organization of this system to the striatal patch and matrix compartments. Coronal sections at three levels through the striatum (A,B,C) are depicted to show the innervation of the patch and matrix compartments from different subsets of midbrain dopamine neurons from three levels (D,E,F). Neurons providing inputs to the striatal matrix compartment (white in D,E,F) are located in the ventral tegmental area (VTA, A10 DA cell group), in the dorsal tier of the substantia nigra pars compacta (in D: SNc, A9) and in the retrorubral area (in F: RR, A8 DA cell group). Neurons providing input to the striatal patch compartment are located in the ventral tier of the substantia nigra pars compacta (in D,E,F: SNc, A9 DA cell group) and from A9 DA cells located in the substantia nigra pars reticulata (in E and F). There is a general topography in that medially located cells project to the ventral striatum and laterally located cells project to the dorsal striatum. Neurons at each rostral-caudal level in the midbrain project rather extensively to throughout the rostral-caudal extent of the striatum such that neurons at level D project to levels A, B and C in the striatum. d e n s e l y p a c k e d , a n d t h e p a r s r e t i c u l a t a , w h i c h is relatively cell sparse c o m p a r e d to the p a r s c o m p a c t a , a n d the r e t r o r u b r a l area, w h i c h lies c a u d a l a n d d o r s a l to t h e s u b s t a n t i a n i g r a ( G e r f e n et al. 1987). T h e d e s i g n a t i o n o f the s u b g r o u p i n g s o f d o p a m i n e n e u r o n s

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according to regional location, A10 cell group in the ventral tegmental area, A9 cell group in the substantia nigra, and A8 cell group in the retrorubral area, conforms to some extent with their projection targets. The A10 dopamine cell group is generally regarded to project to limbic forebrain areas, such as the septal area, prefrontal cortex, olfactory tubercle and nucleus accumbens. The A9 and A8 cell groups are generally regarded as the origin of the projection to the striatum. As is evident, dopamine containing neurons, which project to the striatum, are distributed in each of these groups, including the A10 cell group, due to the inclusion of the nucleus accumbens within the striatum. As is also seen, these neurons are distributed in a somewhat continuous manner, such that delineation of subgroupings based regional location is somewhat arbitrary. A different parcellation of these neurons is suggested based on the morphology of neuronal dendrites, the expression of the calcium binding protein calbindin, and their projection to either the patch or matrix striatal compartments (Gerfen et al. 1987a,b). Using these determinants the projection of midbrain dopamine neurons to the striatum reveals the following organization. Two sets of striatal projecting dopamine neurons are distinguished, a dorsal and ventral tier. The dorsal tier set provides inputs to the striatal matrix compartment. This set encompasses a continuous group which includes those dopamine neurons projecting to the striatum in the ventral tegmetnal area, the dorsal part of the substantia nigra pars compacta, and the retrorubral area. Several other characteristics apply to this set. First, those neurons in the pars compacta are distinguished by the extension of dendrites within the plane of the pars compacta, distinguished from those of the ventral tier. Second, most of the dorsal tier neurons express, in addition to dopamine, the calcium binding protein, calbindin. Third, there is a rough topography to the organization of the projections to the striatum such that more medially situated neurons project ventrally to the nucleus accumbens and ventral striatum, whereas more lateral and caudal neurons, in the A9 and A8 cell groups, project to the dorsal striatum. The ventral tier set provides inputs to the striatal patch compartment. Neurons in this set are situated in the ventral part of the substantia nigra pars compacta and in groups of cells embedded in the pars reticulata. Ventral tier pars compacata neurons are distinguished by their extension of dendrites ventrally into the pars reticulata. Ventral tier dopamine neurons do not display calbindin immunoreactivity. These neurons display a topographic organization in their projections to the striatum, with dorsally positioned neurons projecting to the patch compartment in the ventral striatum and nucleus accumbens, and ventrally positioned neurons, in the pars reticulta projecting to the dorsal striatal patch compartment. It is worthwhile to note that the numbers of dopamine neurons located in the ventral substantia nigra pars reticulata increases at more caudal levels. Consequently, the common view of the substantia nigra as being composed of two separate zones, a dorsal pars compacta in which dopamine neurons are located, and a ventral pars reticulata in which GABA neurons are located, applies only to the rostral most levels of this nucleus. This organization appears to be common across species from rat to primates. 10.2. STRIATAL OUTPUTS The basis of striatal patch-matrix organization is the segregation of striatal medium spiny neurons that have projections to different components of the substantia nigra and entopeduncular nucleus (Gerfen 1984, 1985; Gerfen et al. 1985). Neurons in the patch compartment project to the location of the ventral tier dopamine neurons, whereas neurons in the matrix compartment project to the location of GABA neurons in the 429

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substantia nigra pars reticulata (Gerfen 1984, 1985; Gerfen et al. 1985). This organization appears common throughout all regions of the striatum, including the ventral striatum and nucleus accumbens. Several lines of experimental evidence have revealed this organization. First, retrograde axonal tracers injected into the dopamine cell rich substantia nigra pars compacta or into the pars reticulata selectively label either patch or matrix neurons (Gerfen 1984; Gerfen et al. 1985). However, these methods are limited by the uncertainty of defining the exact area of uptake of transported tracer. Second, the same result has been obtained in several species, including rats, cats and primates (Gerfen 1984, 1985; Gerfen et al. 1985; Jimenez and Graybiel 1989). Third, the calcium binding protein calbindin selectively labels striatonigral neurons in the matrix compartment and not in the patches (Gerfen et al. 1985). Calbindin immunoreactivity is also contained in the terminals of striatal matrix axon projections to the substantia nigra. The distribution of such terminals is concentrated in the pars reticulata and is absent in both the area in which dopamine neurons are located in both the pars compacta and in those parts of the pars reticulata in which dopamine neurons are located. This distribution pattern confirms axonal tracing studies which suggest that patch compartment neurons projections target dopamine neurons in the substantia nigra and that matrix compartment neurons provide inputs to the GABA neurons in the substantia nigra pars reticulata. A parallel organization appears to also apply to the striatal projection to the entopeduncular nucleus (Rajakumar et al. 1993). The entopeduncular nucleus, the rodent homologue of the internal segment of the internal segment of the globus pallidus in primates, may be considered to be part of a continuous group of GABA neurons that extend into the substantia nigra pars reticulata and provide the major output of the basal ganglia. Similar to the GABA neurons in the pars reticulata, entopeduncular neurons provide a projection to the thalamus (Van der Kooy and Carter 1981).. The thalamic targets of the entopeduncular nucleus, and the internal segment of the globus pallidus in primates, provide projections to frontal cortical areas involved with axial musculature. This is contrasted with thalamic targets of the output of the substantia nigra pars reticulata, which are nuclei that provide inputs to frontal cortical areas involved with eye and head movements. Thus the entopeduncular nucleus and substantia nigra pars reticulata appear to form a continuous somatotopically organized output of the basal ganglia. The entopeduncular nucleus is also distinct from the substantia nigra in lacking an associated dopamine cell group. However, like the substantia nigra the entopeduncular nucleus may be divided into two parts on the basis of output neurons. In addition to those entopeduncular nucleus neurons that project to the ventral lateral thalamus, there is a medially situated part of the nucleus which provides inputs directed to the lateral habenula (Van der Kooy and Carter 1981). The lateral habenula in turn projects to the substantia nigra pars compacta, as well as to brain stem nuclei including the medial and dorsal raphe and to the midbrain tegmentum. Recent studies have shown that the striatal matrix compartment provides inputs directed to the thalamic projecting part of the entopeduncular nucleus, whereas the patch compartment provides inputs to the habenular projecting part of the nucleus (Rajakumar et al. 1993). Given the connections through the lateral habenula to the substantia nigra pars compacta, the organization of the patch-matrix projections to the entopeduncular nucleus suggest that a similar, though different, organization as that seen to the substantia nigra from these striatal compartments. Of note is the fact that whereas the direct striatal patch projection to the substantia nigra pars compacta 430

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appears to target the ventral tier dopamine cell group, the patch projection system through the entopeduncular-habenular connections appears to provide inputs directed to the dorsal dopamine cell group. As discussed above the dorsal dopamine cell group provides input directed to the matrix compartment, whereas the ventral dopamine cell group provides input directed to the patch compartment. One missing piece of connectional data concerning the organization of the patch and matrix compartments is the identification of striatal patch neurons that project to the globus pallidus. Whereas the projection of patch neurons to the substantia nigra and entopeduncular nucleus have been described, these neurons account for only half of the projection neurons in the patch compartment. The other half provide inputs to the globus pallidus (Gerfen and Young 1988). One possibility might have been that patch and matrix neurons provide differential inputs to the striato-pallidal border region versus the central region of the globus pallidus, as these regions are distinguished by the dendritic organization of pallidal neurons and by the existence of dual projections from the striatum. However, it has been clearly demonsrated that individual neurons in the striatum provide axon collaterals to both pallidal regions (Chang et al. 1981; Kawaguchi et al. 1990). Another possibility is that patch neurons might provide a select input to cholinergic neurons in the globus pallidus. These neurons, which are scattered in the dorsal globus pallidus and more numerous in the ventral pallidum, have been shown to receive synaptic input from the striatum (Grove et al. 1986). However, it remains purely speculative whether such inputs originate within the striatal patch compartment. Such a connection makes functional sense in terms of the symmetry of the system, but remains to be examined. The segregation of medium spiny neurons with different projection targets to the patch and matrix compartments provides a morphologic basis for these compartments. Moreover, the dendrites of medium spiny neurons appear to remain confined to the compartment of the parent neuron. This has been established with retrograde axonal tracing studies (Gerfen et al. 1985), with Golgi impregnation studies (Bolam et al. 1988) and most directly with intracellular labeling of individual neurons (Kawaguchi et al. 1989). These latter studies have demonstrated that the dendrites may take tortuous paths to remain confined within a particular compartment. In particular, patch neurons are often seen to have recurved dendrites that dutifully respect the borders between the patch and matrix compartments. This organization of the dendrites of medium spiny neurons suggests that afferents from outside the striatum that target a particular compartment 10.3. CORTICAL INPUTS Cortical inputs to the patch and matrix compartments originate in different sublayers of layer 5, from most cortical areas (Gerfen 1989). Cortical inputs were amongst the first to be described as being compartmentally organized. Initial studies suggested that cortical areas with limbic connections provide inputs directed to the patch compartment, whereas neocortical areas provide inputs to the matrix compartment (Donoghue and Herkenham 1986; Gerfen 1984). However, more detailed analysis revealed that although different cortical areas provide inputs that differ in magnitude to the two compartments, most cortical areas appear to provide inputs to both compartments (Gerfen 1989). For example, the prelimbic cortex in the rat, which is on the medial bank of the frontal cortical hemisphere, provides a dense input to the patch compartment in the medial striatum from neurons in the deep part of layer 5 and an input to the matrix compart431

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E I= 1 mm Fig. 26. The relationship between the laminar organization of the cortex and the striatal patch-matrix compartments is diagrammed showing inputs from the prelimbic (A) and cingulate (D) cortices to the striatum. Corticostriatal neurons located in the deeper part of layer 5 (black cells in A and B) in each cortical area provide inputs directed to the patch compartment (black stippling in B,C, E and F), whereas corticostriatal neurons located in the superficial part of layer 5 (white cells in A and D) provide inputs directed to the matrix compartment (white stippling in B,C, E and F). Inputs from these cortical areas are somewhat greater to the patch compartment as compared to their inputs to the matrix compartment. The prelimbic and cingulate cortical areas provide inputs to a topographically related region in the striatum which overlaps to some extent for these two corticostriatal projections.

ment surrounding these patches from the superficial part of layer 5. This organization has subsequently been confirmed to also apply to cortical inputs to the ventral striatum and nucleus accumbens (Berendse et al. 1992a). The determination of the compartmental target of neurons in different sublaminae of the prelimbic cortex was based on a large number of cases of injections of PHA-L 432

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Fig. 27. The relationship between the laminar organization of the cortex and the striatal patch-matrix compartments is diagrammed showing inputs from the medial agranular cortex (AGm: A) and lateral agranular cortex (AGI: D) to the striatum. Corticostriatal neurons located in the deeper part of layer 5 (black cells in A and B) in each cortical area provide inputs directed to the patch compartment (black stippling in B,C, E and F), whereas corticostriatal neurons located in the superficial part of layer 5 (white cells in A and D) provide inputs directed to the matrix compartment (white stippling in B,C, E and F). Inputs from these cortical areas are relatively greater to the matrix as compared to the patch compartment. Also of note is the discontinuous pattern of inputs that arise from these injections to the striatum.

into a specified cortical area, such as the prelimbic area (Gerfen 1989). In order to assure that an injection was confined to the prelimbic cortex, that is, labeled cortical neurons whose efferent axons were labeled by the tracer, and did not include injected neurons in adjacent cortical areas several criteria were applied. The first was by inspection of the injection site to determine that labeled neurons were confined within a single cortical 433

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area. The second criterion was the pattern of thalamic labeling. Thus, the pattern of thalamic labeling of injections into the prelimbic area was compared with injections into adjacent cortical areas. Cases were selected for inclusion in a set of injections only if the pattern of thalamic labeling could be distinguished between that of the anterior cingulate and medial agranular cortices, the areas adjacent to the prelimbic cortex. The third criterion was that the pattern of the crossed cortico-cortical labeling displayed a pattern in which the crossed projection system was concentrated over the contralateral homologous cortical area. Using these criteria it was found that injections that were confined to a single cortical area such as the prelimbic area could be grouped into three types, those with projections to the striatal patch compartment, those with projections concentrated in the striatal matrix compartment and those with projections to both compartments. In these experiments the patch and matrix compartments were identified with either naloxone binding or calbindin immunoreactivity. In all types the area of the striatum innervated was comparable, although small differences reflecting microtopographic organization were apparent. Several features of the patterns of cortical labeling distinguished the injections which labeled projections to the patch compartment as compared to those which labeled inputs to the matrix compartment. Injections that labeled inputs preferentially distributed in the patch compartment labeled neurons that were situated in deeper parts of layer 5. In addition, the labeling of axon collaterals of these labeled neurons was distributed in the prelimbic area in layer 5 and 6 with little labeling of axons in superficial layers. A comparable pattern of labeling was also observed in the contralateral homologous cortical area. Contrasted with this pattern of labeling was that of injections which labeled inputs directed preferentially to the matrix compartment. In these cases labeled neurons were located more superficially, in upper layer 5 and also in layers 2 and 3. The pattern of axonal labeling surrounding the injection site showed dense labeling in superficial cortical areas. A comparably dense distribution of labeled fibers was also observed in the superficial layers of the contralateral cortex. While interpretations are limited by methodological considerations, the pattern of labeled projections suggested that neurons projecting to the striatal patch compartment are located more deeply in the cortex than those which provide projections to the matrix compartment. A similar organization was also found with injections into the infralimbic, anterior cingulate, medial agranular and lateral agranular cortices (Berendse et al. 1992a; Gerfen 1989). In each of these other cortical areas, the same criteria were applied for assuring that injections were confined to a single cortical area. In addition, the same pattern of cortical labeling was also observed. Differences between these cortical areas were mainly related to the relative density of inputs to the patch compartment. Infralimbic and prelimbic cortical injections provided denser inputs to the patch compartment, while cingulate and medial agranular cortical areas showed a slightly less dense input to the patch compartment. Inputs from lateral agranular cortical injections showed only very sparse inputs to the patch compartment. In addition, projections from the prelimbic, infralimbic, and cingulate cortices were each distributed to a rather continuous region within the striatum, including both the patch and matrix compartments. On the other hand, the pattern of striatal labeling after injections into the lateral agranular cortex were decidedly discontinuous, with separated dense zones of labeling within the matrix. Of note is the fact that the patterns of cortical labeling from such cortical injections also show marked discontinuities. This aspect of cortico-cortical and cortico-striatal labeling is of interest in the context of the relationship between cortico-cortical and corticostriatal organization. 434

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Thus, it appears that many, and perhaps most, cortical areas provide inputs to both the patch and matrix compartments (Gerfen 1989). However, the relative density of inputs to the patch compartment is denser from periallocortical areas such as the infralimbic and prelimbic cortices. Conversely, neocortical areas such as the medial and lateral agranular cortices have a relatively greater input to the matrix as compared to the patch compartment. Differences in the relative inputs from different cortical areas have led to the suggestion that inputs from the cortex to the patch-matrix compartments is related to the cortical area of origin. In this view cortical areas may be viewed as a continuum of areas with inputs directed to the patch compartment from allo- and peri-allocortical areas and those with inputs to the matrix compartment from neocortical areas. However, such a view should not be confused with the realization that the inputs from a given cortical area are directed to both compartments, albeit to different extents. Studies in primates have revealed a predominance of inputs to the matrix compartment. However, most studies have examined inputs from neocortical areas, which would be predicted to have a relatively weak input to the patch compartment. Recent studies have reported inputs to the patch compartments from primate cortical areas comparable to those in rats which also have a predominant input to the patch compartment. This would suggest that a similar organization applies in primates as well. 10.4. THALAMIC AFFERENTS Thalamic afferents in the striatum from the parafascicular/intralaminar nuclei are organized relative to the patch matrix compartments (Beckstead 1985; Berendse et al. 1988; Herkenham and Pert 1981; Xu et al. 1991). Inputs from the parafascicular/ centromedian thalamic nuclei provide inputs directed to the matrix compartment. Inputs to the striatal patch compartment arise from more restricted parts of the intralaminar thalamic nuclei. 10.5. GENERAL PATCH-MATRIX ORGANIZATION The general organization of the patch-matrix compartments provides separate pathways from the cortex, through the striatum to differentially effect dopamine and other, basal ganglia feed-back circuits, or to affect basal ganglia GABAergic output neurons in the entopeduncular nucleus and substantia nigra pars reticulata. Thus, the cortical connections through the patch compartment appear to be related to regulation of the dopamine, and possibly serotonergic feedback systems to the striatum, whereas cortical connections through the matrix compartment appear to be related to regulation of the output neurons of the basal ganglia. This organization appears to be common to all parts of the striatum. There has been some confusion in the literature with suggestions that the cortical inputs and striatal outputs of the patch-matrix compartments in the ventral striatum differ from those in the dorsal striatum. However, the differences that have been suggested are related to the use of markers for identifying patch-matrix compartments. Some studies of the ventral striatal patch-matrix organization have used enkephalin immunoreactivity as a compartmental marker (Berendse et al. 1992a, 1992b). However, this marker shows a transition between the dorsal and ventral striatum that shifts relative to other neurochemical markers such as calbindin and opiate receptor binding (Voorn et al. 1989), which are more consistent with connectional definitions of patch-matrix compartmental 435

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organization (Gerfen et al. 1985). When patch-matrix compartments are defined on the basis of input-output connectional organization the organization in the dorsal and ventral striatum is identical. 436

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10.6. CORTICAL ORGANIZATION RELATED TO STRIATAL PATCH-MATRIX COMPARTMENTS The relationship between the cortical laminar organization and striatal patch-matrix compartments suggests that the cortical output systems may be organized to regulate the dopamine feedback system to the striatum, which in turn regulates the cortical projection through the matrix compartment to the basal ganglia output neurons. This concept was originally formulated in terms of limbic cortical inputs to the patch compartment (Gerfen 1984). However, with the finding that all cortical areas may provide inputs to the patch compartment (Gerfen 1989), albeit to varying extents, the idea of a preferential connection between 'limbic' cortices and the patch compartment needs to be reexamined. We have suggested that the relationship between the cortex and the striatal patch-matrix compartments is related to the laminar organization of the cortex. This then raises the question as to the functional organization of cortical lamination. Neurons in the cortex are segregated into laminae in which neurons with similar projections are grouped (Jones 1984). For example, pyramidal neurons in superficial layers 2 and 3 provide axonal connections within the cortex, pyramidal neurons in layer 6 are the source of projections to the thalamus, and pyramidal neurons in layer 5 provide other subcortical projections, including those to the striatum. These patterns of cortical efferents related to the cortical layer of origin are by no means absolute. Moreover, there is considerable heterogeneity amongst the classes of cortical projections from a given layer. Relevant to the topic of this discussion are the different types of neurons in layer 5 and particularly the different types of corticostriatal neurons. As described above there are at least 3 types of corticostriatal neurons, pyramidal tract neurons with collaterals into the striatum, cortico-thalamic neurons with collaterals into the striatum, and bilaterally projecting cortico-striatal neurons. One possibility is that these different types of corticostriatal neurons contribute differentially to the projections to the patch and matrix compartments. However, at this time there is not sufficient data to determine whether such a distinction exists. Another possibility is that different subsets of each of these different corticostriatal types project to both compartments. The question therefore remains open as to what might distinguish cortical neurons that project to the striatal patch and matrix compartments. One of the determining features of PHA-L injections into the cortex which distinguished patch from matrix projections was the pattern of cortico-cortical projections of the injected neuron population. Injections which selectively labeled inputs to the patch compartment labeled both local and contralateral cortico-cortical connections that were preferentially distributed in deeper cortical layers. Conversely, injections which selectively labeled inputs to the matrix compartment labeled both local and contralateral cortico-cortical connections that were preferentially distributed to superficial cortical layers. With this method it is not possible to attribute the cortico-cortical pattern of labeling to neurons that project also Io the striatum. However, it is possible to speculate that a difference in the corticocortical connections of patch and matrix projecting corticostriatal neurons might distinguish these two neuron types. There do exist layer 5 neurons that show such patterns of cortico-cortical connectivity. For example, at least two distinct types of layer 5 neurons have been distinguished on the basis of their local axon collateras, one type which has axon collaterals which distribute longitudinally in layer 5 and 6, whereas another neuron type has axon collaterals that distribute to superficial layers, 2 and 3 (Chagnac-Amitai et al. 1990). These latter axon collaterals display a much more restricted distribution in the longitudinal axes. If such differences in local cortical axon 437

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collaterals apply to patch- and matrix-directed corticostriatal neurons the implication is that patch-directed corticostriatal neurons influence a larger domain of the cortical area of origin than do matrix-directed corticostriatal neurons. The resolution of this speculation awaits single cell labeling analysis. A possible function of the laminar organization of the cortex and the patch-matrix compartmentation of the striatum might be inferred from the apparent transition in the relative contribution to the striatal compartments from allo- (or periallo-) cortical compared to neocortical areas. As discussed above, early studies had suggested a preferential input from cortical areas connected with the limbic system to the striatal patch compartment, which in turn provides a direct input to dopamine neurons that project back to the striatum. This concept was a modification of the ideas put forward by Nauta and his co-workers that the basal ganglia was a site of integration of limbic and non-limbic systems. They suggested that the limbic parts of the striatum, the ventral striatum including the nucleus accumbens, which is the target of 'limbic' inputs from the amygdala, hippocampus, and olfactory related cortical areas, provided the main input to the dopamine neurons in the substantia nigra pars compacta, which projected back to both ventral, 'limbic'- and also dorsal, non-limbic-striatal regions. Initial studies of the input-output organization of the patch-matrix compartments, modified the ideas of Nauta with the finding that it was the patch compartment neurons, in both the ventral and dorsal striatum, which are the source of inputs to the dopamine feedback neurons to the striatum. Our early analysis retained the concept of Nauta, by suggesting that the source of the input to the patch compartment was exclusively from limbic connected cortical areas. This idea was further modified with more recent findings that the source of inputs to the patch compartment was not restricted only to limbic-connected cortical areas. Thus, given the fact that the patches in the dorsal striatum receive inputs from neocortical, non-limbic, areas of cortex, and that these patches nonetheless provide inputs to the dopamine nigrostriatal feedback system requires a further modification of the concept of the integration of so-called 'limbic' and 'non-limbic' integration within the basal ganglia. Rather than consider that the striatal patch-matrix systems are related to limbic and non-limbic cortical areas defined on the basis of the former's connections with olfactory related structures, these cortical areas might be considered on the basis of the differences in their organization. For this discussion it is best to refer to allo- and peri-allocortical and neocortical areas because of the obvious conceptual difficulties of defining limbic and non-limbic cortices. Allo-, peri-allo- and neo-cortical areas appear to process information in distinct ways. For example, in the piriform cortex, which may be viewed as a prototypic allocortical area, information coding of specific odors is distributed throughout the cortical area. Different odors are encoded in the same cortical space such that specific odors are encoded by different patterns of activity distributed across that cortical area. On the other hand, in neocortical areas such as the somatosensory or motor cortex, the encoding of information is somatotopically organized within the cortical fields. The computational differences required for encoding these different modalities, olfaction on the one hand, and somatotopically organized information on the other, is reflected in the organization of cortico-cortical connections within each of these cortical areas. Thus in the piriform cortex each neuron is broadly connected with other neurons in this cortical area, whereas in neocortical areas there appears to be considerable local specificity of cortico-cortical connections. Thus, allocortical areas appear to process information with relatively uniformly distributed connectional patterns, whereas neocortical areas parse information on the basis of somatotopic or retinotopic maps. Alheid and Heimer (1988) have also proposed that the subcortical 438

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connections of allo- and neocortical areas share common general organizational schemes whose specific elements reflect a transition in the final targets of these systems and in the feedback mechanisms they employ. In general they suggest that the projections of allo-cortical areas target more direct subcortical feedback systems, whereas neocortical areas target indirect subcortical feedback systems that are organized to provide more specific feedback to the cortex through the thalamus. We would argue that the transition from distributed information processing typical of allocortex to spatially compartmentalized information processing typical of neocortex is relevant to the function of the striatal patch-matrix compartments. The patch compartment, appears to have connections analogous to those of allocortex, providing inputs to a more direct feedback system, the nigrostriatal dopamine system. Conversely, the matrix compartment, appears to have connections analogous to the neocortex, which target indirect feedback pathways to the cortex through the thalamus. Whereas the connections of allocortical areas and somatosensory and visual cortical areas represent the extremes of the two forms of information processing, most cortical areas encompass both schemes, but to varying degrees. Thus, it is suggested that the the striatal targets of allocortical areas, which may be the shell region of the nucleus accumbens, whereas, the striata| target of the most extreme neocortical areas, which target striatal areas that are relatively devoid of the patch compartment, represent extreme cases. In most of the striatum both patch and matrix compartments exist for these two types of information processing. This would suggest that there is a retention of some organizational elements of allocortex in the transition to neocortex. It is proposed from all cortical areas providing inputs to both compartments, that cortical neurons projecting to the patch compartment have allocortical type connectional features, whereas those projecting to the matrix have 'neocortical' type connectional features. The relative numbers of each type varies according to the cortical area of origin. Regardless of the speculative proposals for the functional significance of the relationship between the cortex and the patch-matrix compartments several determinants of this relationship may be stated with some certainty. First, there is a differential projection of different cortical neurons to the patch and matrix compartments. Second, both patch and matrix corticostriatal projection neurons are located within a single cortical area. A corollary of this is that most cortical areas appear to contain both patch and matrix projection neurons but the relative number of each varies according to the type of cortical area. Third, within a single cortical area patch and matrix corticostriatal neurons are preferentially located in different sublaminae. As laminar organization varies across cortical areas the precise distribution of the patch and matrix corticostriatal projecting neurons may also vary. The laminar organization of patch-matrix corticostriatal neurons is distinguished from other aspects of cortical organization, such as the columnar organization. Fourth, the organization of separate cortical pathways into the patch and matrix compartments are carried through the striatum to provide segregated inputs to dopamine neurons and GABA output neurons in the substantia nigra.

11. DIRECT/INDIRECT STRIATAL OUTPUT SYSTEMS 11.1. C O N N E C T I O N A L BASIS Medium spiny neurons have a common morphology in terms of their size, dendritic organization and local axon collaterals. Within the striatum these neurons have an axon 439

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Fig. 29. Coronal sections through the striatum showing mu-opiate receptor with 3H-naloxone binding of patches (A and B) and in adjacent sections spiny projection neurons labeled by in situ hybridization histochemistry with probes directed against substance P mRNA (A') and enkephalin mRNA (B'). Substance P and enkephalin are expressed by different populations of spiny projection neurons, each comprising about half of the population and each evenly distributed in both patch and matrix compartments (arrows show patches in the corresponding sections). From Gerfen and Young (1987).

collateral that extends within varying domains around the parent neuron. Each of these neurons provides an axon that projects out of the striatum. Studies in which individual striatal medium spiny neurons were filled with the marker biocytin revealed subsets of neurons on the basis of the projection axons (Kawaguchi et al. 1990). One type, referred to as a striatopallidal neuron, provides an axon that extends into the globus pallidus and arborizes extensively, usually in two separate domains within this nucleus. These neurons do not have an axon that extends beyond the globus pallidus. A second type extends an axon collateral into the globus pallidus, which does not arborize extensively, and extends other collaterals that extend into either the entopeduncular nucleus and/or the substantia nigra. In order to simplify the terminology this second type of neuron is 440

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ENK

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Fig. 30. In situ hybridization histochemical localization of mRNAs to identify peptides and dopamine receptor subtypes in striatal spiny projection neurons. Striatonigral neurons contain both D 1 and substance P-mRNAs, whereas striatopallidal neurons contain both D2 and enkephalin mRNAs. A-C) Neurons that project to the substantia nigra have been retrogradely labeled with the fluorescent dye fluorogold (whitish labeled cell bodies). In situ hybridization labeling of specific mRNAs is shown by white grains. A) D 1 dopamine receptor mRNA is localized in labeled striatonigral neurons (arrows). B) Substance P mRNA is also localized in labeled striatonigral neurons(arrows). C) D2 dopamine receptor mRNA is not contained in labeled striatonigral neurons but in unlabeled striatopallidal neurons (open arrows). D) Enkephalin mRNA is also contained in unlabeled striatopallidal neurons (open arrows). E) Both D 1 and D2 mRNAs are labeled in the same section, D1 mRNA with an S35-riboprobe that is marked by white silver grains over neurons and D2 mRNA with a digoxigenin-riboprobe that is labeled with a dark immunoreactive reaction. D1 and D2 mRNAs are segregated in separate neurons, with less than 5% of the entire population of striatal spiny projection neurons containing appreciable amounts of both receptor subtypes. A-C) from Gerfen et al. 1990.

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~ntia ra Fig. 31. Summary diagram of the 'direct' and 'indirect' striatal output pathways. Layer 5 cortical neurons provide excitatory input (+) to the striatum. The direct striatal projection is provided by D1/substance P/dynorphin-containing neurons to the substantia nigra and entopeduncular nucleus, and to a lesser degree to the globus pallidus. The indirect striatal projection is provided by D2/enkephalin-containing neurons that project to the globus pallidus. The globus pallidus in turn provides an inhibitory projection to the substantia nigra and to the subthalamic nucleus. The subthalamic nucleus provides an excitatory input to the substantia nigra. Thus, the 'direct' and 'indirect' pathways provide antagonistic input to the substantia nigra. The GABA neurons in the substantia nigra provides an inhibitory projection to the superior colliculus, pedunculopontine nucleus (not shown) and thalamus. The thalamic nuclei receiving this output project back upon the frontal cortex. The entopeduncular nucleus is connected in a similar manner to the substantia nigra but is not shown.

442

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referred to as a striatonigral neuron. This simplification is used for several reasons, but the fact that it is a simplification should be kept in mind when considering the functional significance of these projection systems. First, as demonstrated, these neurons do extend an axon into the globus pallidus. Although the extent of arborization of this axon collateral is less than that of the striatopallidal neuron, it exists, and may make functional synapses with pallidal neurons. The numbers of neurons so contacted and the relative input of this neuron relative to that of the striatopallidal neuron input is not yet known. However, in terms of the numbers of synapses, striatonigral axon collaterals in the globus pallidus make as many as half the number of synaptic contacts as the axons of striatopallidal neurons. Second, the entopeduncular nucleus (as well as the internal segment of the globus pallidus in primates) and the substantia nigra may be considered to be part of a single nuclear complex in terms of both their inputs and outputs. Both structures receive direct inputs from the striatum and both contain GABA neurons that may be considered to be part of the output system of the basal ganglia in that they project to the thalamus. The targets in the thalamus are distinct (Gerfen et al. 1982; Van der Kooy and Carter 1981). The entopeduncular nucleus projects to the ventral lateral thalamic nucleus and lateral habenula, whereas the substantia nigra pars reticulata provides inputs to the ventral medial and intralaminar thalamus. These different targets of the two output components of the basal ganglia reflect that topographic organization of striatal outputs. Intracellular staining studies confirmed prior retrograde tracing experiments, in which tracers were injected into the target nuclei, which demonstrated that striatal neurons projected to either the globus pallidus or substantia nigra (Beckstead and Cruz 1986; Gerfen and Young 1988; Loopuijt and Kooy 1985). Such studies established several features of the striatal organization of such neurons. First, there appear to be distinct neuron subpopulations. In addition to the direct demonstration of this from Kawaguchi's work, this is also suggested from retrograde studies. Studies employing two fluorescent markers, one injected into the substantia nigra and the other into the globus pallidus, have shown most neurons to be labeled from only one injection site (Beckstead and Cruz 1986; Loopuijt and Kooy 1985). Even accounting for the possibility of injections that are not perfectly matched topographically in the two structures, such a labeling pattern points to an inherent limitation of the technique, as, based on the existence of a collateral of striatonigral nuerons in the globus pallidus the pattern of labeling is almost certainly revealing the precise organization of axonal projections. In this case such a limitation is an asset in that it does reveal two connectionally distinct neuron types. Second, the numbers of each projection type appear to be approximately equal. Given that over 90% of striatal neurons are projection neurons, estimates based on retrograde labeling suggest that approximately 40-45% project principally to the globus pallidus and another 40-45% project principally to the substantia nigra. Third, striatopallidal and striatonigral neurons are interspersed with one another. In some cases they may form small clusters of 2-5 neurons projecting to one site. In other cases, neurons projecting to separate sites may be nearest neighbors, often appearing to be in close apposition. 11.2. PEPTIDE BASIS The segregation of striatal output neurons on the basis of their differential targetting of the globus pallidus and substantia nigra was first suggested by the immunohistochemical labeling of opiate and tachykinin peptides in striatal terminals in these nuclei. 443

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Fig. 32. Data from an experiment demonstrating D 1- and D2-receptor selective gene regulation in 'direct' and 'indirect' striatal projection neurons. Images are of coronal sections of film autroadiographs labeled with in situ hybridization histochemical localization of enkephalin mRNA (top row), substance P mRNA (middle row) and dynorphin mRNA (bottom row) from an intact rat striatum (control striatum: first column), from a striatum depleted of dopamine (6-OHDA lesion: second column), from a dopamine depleted striatum of an animal treated with the D1 agonist SKF38393 (single daily injections 5 mg/kg for 21 days), and from a dopamine depleted striatum of an animal treated with a D2 agonist quinpirole (continuous treatment of lmg/kg for 21 days). Dopamine depletion elevates enkephalin, decreases substance P and has little effect on dynorphin. Subsequent D 1 agonist treatment has no effect on enkephalin (contained in D2 bearing neurons) but reverses the lesion induced decrease in substance P and causes a large increase in dynorphin mRNA both of which are contained in D 1 bearing neurons. On the other hand, subsequent to dopamine depletion of the striatum, D2 agonist treatment reverses the lesion-induced elevation of enkephalin mRNA in neurons bearing D2 receptors, but has no effect on substance P or dynorphin in D1 bearing neurons. From Gerfen et al. 1990.

Striatal neurons projection neurons all contain GAD (Aronin et al. 1984; Kita and Kitai 1988; Ribak et al. 1979), although subpopulations contain different neuropeptides including the opiate peptides enkephalin (Beckstead 1985; DiFiglia et al. 1982; Haber and Watson 1983; H6kfelt et al. 1977; Pickel et al. 1980) and dynorphin (Vincent et al. 1982a; Vincent et al. 1982b), or the tachykinin substance P (Brownstein et al. 1977; Hong et 444

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A diagram of the model of Parkinson's disease that suggests that dopamine (DA) depletion in the disease results in an elevated output of the indirect pathway (enkephalin, ENK, D2 containing neurons), which results in increased excitatory input from the subthalamic nucleus (stn) to the internal globus pallidus (GPi) and substantia nigra and a decreased output of the direct pathway (substance P: SP, D1 containing neurons). See text for further details.

al. 1977; Kanazawa et al. 1977). Immunohistochemical studies showed that these peptides are localized in connectionally defined striatal output neurons (Beckstead and Kersey 1985; Haber and Watson 1983). Enkephalin-immunoreactive terminals, originating from axons of striatal neurons, are concentrated in the globus pallidus, with only sparse distributions in the substantia nigra pars compacta (Beckstead and Kersey 1985; Haber and Watson 1983). Conversely, both dynorphin and substance P show dense terminal immunoreactivity in the substantia nigra (and entopeduncular nucleus), and only a sparse distribution in the globus pallidus (Brownstein et al. 1977; Hong et al. 1977; Kanazawa et al. 1977; Vincent et al. 1982b).. Whereas such studies had established 445

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the striatal origins of the terminal labeling in these structures, early immunohistochemical techniques were unable to identify the cells of origin without the use of colchicine. Moreover, peptide immunoreactivity in the striatum revealed complex patterns of heterogeneity being highly concentrated in the patch compartment (to be discussed in detail later) (Graybiel et al. 1981). However, these patterns varied from region to region which led to some ambiguity concerning the compartmental relationships of the neurons containing the different peptides. In part these patterns of immunohistochemical localization reflect technical aspects of the method in that different fixatives revealed different patterns of labeling (Graybiel and Chesselet 1984). As has become evident the varied levels of peptides in different striatal compartments and in different regions reflects regulatory mechanisms that underlie the functional organization of the striatum (Gerfen 1991). That the relative peptide levels in striatal neurons may be considered distinct from the localization of peptides in connectionally defined striatal neurons is evident from studies that combine axonal tracing techniques with in situ hybridization histochemical localization of the messenger RNAs that encode the various peptides (Gerfen and Young 1988). Using these techniques it has been established that striatopallidal neurons contain mRNA encoding enkephalin and striatonigral neurons contain mRNAs encoding both dynorphin and substance P. Moreover, these studies show that these two connectionally defined neuron types each constitute approximately half of the striatal projection neuron population, that the two populations are intermingled with each other throughout all regions of the striatum and that they are equally distributed in both the patch and matrix compartments. This pattern of localization is shown in Figures 29 and 30. 446

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11.3. DOPAMINE RECEPTOR-MEDIATED REGULATION Levels of peptides in striatal neurons are regulated by dopamine receptor-mediated mechanisms. Work by several groups demonstrated that levels of enkephalin and substance P are oppositely modulated by dopamine (Gerfen et al. 1991; Hong et al. 1978a; Hong et al. 1978b; Young et al. 1986). Dopamine depletion of the striatum or neuroleptic blockade of D2 dopamine receptors result in an elevation of enkephalin peptide and mRNA levels in striatopallidal neurons (Hong et al. 1978b, 1985; Mocchetti et al. 1985; Tang et al. 1983) and a decrease in substance P levels (Bannon et al. 1986; Hanson et al. 1981; Hong et al. 1978a). Conversely, pharmacologic treatments that enhance dopamine neurotransmission result in elevated substance P and dynorphin peptide and mRNA levels in striatonigral neurons (Gerfen et al. 1990, 1991; Hanson et al. 1987; Li et al. 1986, 1988). The opposite effects that dopamine has on the peptides in striatal output neurons appear to be related to the differential expression of the D 1 and D2 dopamine receptor subtypes by the neurons that express these peptides (Gerfen et al. 1990). Thus, mRNA encoding the D 1 dopamine receptor subtype is localized in striatonigral neurons and the mRNA encoding the D2 dopamine receptor subtype is localized in striatopallidal neurons. This distribution of receptors has been established in numerous ways. First, similar to the immunohistochemical localization of peptides to the different output pathways, receptor binding studies demonstrate a differential localization of D2 and D 1 receptor binding, of striatal origin, in terminals in the globus pallidus and substantia nigra, respectively (Beckstead 1988; Richfield et al. 1989). Second, combined axonal tracing and in situ hybridization studies localize D2 and enkephalin m R N A in striatopallidal neurons and D1 and substance P m R N A in striatonigral neurons (Gerfen et al. 1990). Third, double in situ hybridization studies reveal the exclusive co-localization of D2 mRNA in neurons with enkephalin mRNA and D1 mRNA in neurons with substance P mRNA (Le Moine et al. 1990, 1991). Fourth, dual visualization of D 1 and D2 mRNAs in the same histologic sections with in situ hybridization demonstrate a near complete segregation of neurons expressing each receptor subtype. It should be noted that, while the in situ hybridization studies seem conclusive when considered alone, they are not in agreement with the results from a number of physiological and biochemical studies, and so the differential expression of D 1 and D2 receptors continues to be controversial. Most striatal spiny neurons, including most identified striatonigral neurons, respond to both D1 and D2 receptor agonists and antagonists, even when isolated from all synaptic input (Surmeier et al. 1992). Similarly D 1 and D2 agonists act synergistically in suppressing Na+-K + ATPase activity in isolated striatal neurons (Bertorello et al. 1990). Gene amplification techniques have been used to demonstrate the co-localization of D1 and D2 dopamine receptor subtype mRNAs in striatonigral neurons (Surmeier et al. 1992). These data are at odds with in situ hybridization techniques. One possiblity is that even though in situ hybridization techniques show a segregation of D 1 and D2 dopamine receptors in different striatal neurons, there may be low levels of expression of D1 mRNA in striatopallidal neurons that express relatively high levels of D2 m R N A and conversely there may be low levels of expression of D2 m R N A in striatonigral neurons that express relatively high levels of expression of D 1 mRNA. The question is whether the disparity of different levels of these receptor subtypes in individual neurons is related to the function of these neurons. Physiologic studies have suggested at least that individual neurons show physiologic changes in ion conductances in response to activation of both receptor subtypes. However, as will be 447

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detailed below studies employing selective D 1 and D2 receptor agonists and antagonists suggest that striatopallidal and striatonigral output neurons are differentially and selectively affected by pharmacological manipulation of these dopamine receptor subtypes when changes in gene regulation are measured (Dragunow et al. 1990; Gerfen et al. 1990; Robertson and Fibiger 1992; Robertson et al. 1990). In animals with 6-hydroxydopamine lesions of the nigrostriatal pathway, it is possible to selectively activate receptor subtypes without the effects of endogenous transmitter (Gerfen et al. 1990, 1991). In such dopamine depleted striata, levels of genes contained in striatopallidal neurons are increased, including mRNAs encoding enkephalin and the D2 receptor. Conversely, in striatonigral neurons various mRNAs are decreased, including those encoding the peptides substance P and dynorphin and the D1 dopamine receptor. These lesion-induced alterations are selectively reversed, in each neuron type, by treatment with agonist directed against the recpetor expressed by that neuron type. Thus, increased enkephalin and D2 receptor mRNA levels are reversed by administration of the D2 agonist quinpirole, and the decreased substance P and D1 receptor mRNA levels are reversed by D1 agonist (SKF38393) treatment. Significantly, the schedule of treatment with these agonists was a critical factor in the effect on peptide mRNA levels. Two treatment schedules were used to administer dopamine receptorselective agonists to animals with unilateral lesions of the nigrostriatal dopamine system. The first was a continuous infusion schedule, in which the drugs were administered for 21 days with osmotic minipumps implanted intraperitoneally. The second was an intermittent schedule, in which drugs were administered once daily for 21 days. Reversal of the lesion induced increase of enkephalin and D2 receptor mRNA was effected with continuous (1 mg/day) but not intermittent (1 x 1 rag/day) quinpirole treatment. Conversely, reversal of the lesion-induced decrease in substance P and D1 receptor mRNA was effected with intermittent (1 x 10 mg/kg) but not continuous (10 mg/day) SKF38393 treatment. In addition, intermittent SKF-38393 treatment resulted in a large increase above baseline levels of the mRNA encoding the peptide dynorphin in striatonigral neurons. These results suggest that gene regulation in striatopallidal and striatonigral neurons are regulated in different ways by the activation of the dopamine receptor subtypes (Gerfen et al. 1990). Changes in peptide/protein or mRNA levels in neurons in response to pharmacologic activiation or blockade of receptor subtypes does not substitute for measurements of physiologic response. Moreover, as changes in peptide levels occur over a prolonged time period, these may be secondary to the direct effect of dopamine receptor activation. However, other markers of gene regulation, such as the induction of transcription factors including the immediate early gene c-fos, which occur immediately following drug treatments, reveal a similar pattern of selective effect of D1 and D2 dopamine receptor subtype effects on striatonigral and striatopallidal neurons. For example, in the unilateral nigrostriatal dopamine lesion model, a single injection of the D1 agonist SKF-38393, results in the rapid induction of c-fos in striatonigral and not in striatopallidal neurons (Robertson et al. 1989, 1990). Thus, the immediate effect of activation of D1 receptor activation appears to have a selective effect on striatonigral neurons. While the dopamine depleted striatum provides a good model for study of the selective effects of D1 and D2 receptor stimulation these effects are abnormal in the sense that the pharmacologic treatments that alter gene regulation in the lesioned striatum are not paralleled in the unlesioned striatum. These differences in effect are not due to a redistribution of the receptor subtypes, as the segregated localization of the D 1 and D2 receptor subtypes to striatonigral and striatopallidal neurons occurs in both the lesioned 448

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and unlesioned striatum (Gerfen et al. 1990; Le Moine et al. 1990, 1991). More likely, such differences reflect altered receptor-mediated signal transduction processes that result in supersensitive responses to receptor activation. However, there are some drug treatments that elicit responses in the normal striatum that are similar to those in the lesioned striatum. For example, systemic administration of the D2 receptor antagonist haloperidol results in the immediate induction of c-fos selectively in striatopallidal neurons (Dragunow et al. 1990; Robertson and Fibiger 1992). Longer-term treatment with such neuroleptics result in elevated enkephalin (Hong et al. 1978b, 1985; Mocchetti et al. 1985; Tang et al. 1983). These effects, in normal striatum, are the converse of changes caused by striatal dopamine depletion and subsequent D2 agonist treatments that selectively effect striatopallidal neurons. In normal rats induction of immediate early genes and changes in peptide levels occur within the striatum after single and repeated administration of drugs that enhance dopamine function. Both amphetamine administration, which acts to enhance dopamine release, and cocaine administration, which acts to prolong the effects of dopamine by blocking catecholamine reuptake, result in c-fos induction in the striatum (Cenci et al. 1992; Graybiel et al. 1990; Steiner and Gerfen 1993; Young et al. 1991). The regional patterns of induction produced by these two drug treatments differ, with amphetamine producing induction that is most prevalent in the striatal patch compartment, whereas cocaine produces induction in both patch and matrix compartments that is regionally localized to the dorsal striatum. In the case of cocaine administration c-fos is induced selectively in striatonigral neurons and this induction is blocked by D1 receptor antagonists (Cenci et al. 1992; Graybiel et al. 1990; Young et al. 1991). These effects provide several insights into dopamine regulation of striatal function. First, they provide evidence that the same underlying dopamine receptor-mediated regulatory processes that occur in the dopamine depleted striatum function in the normal striatum. Second, the compartmental and regional variations in the response of striatal neurons to different manipulations of dopamine function in the striatum suggest heterogeneity in the organization of nigrostriatal dopamine system and other striatal afferent systems, most notably the corticostriatal and thalamostriatal systems. 11.4. OTHER (NON-DOPAMINERGIC) REGULATORY RECEPTOR SYSTEMS IN STRIATUM The organization of D1 ad D2 dopaminergic receptors among the striatonigral and striatopallidal neuron populations is relatively uniform throughout all regions of the striatum. Moreover, the opposite regulation of these two populations of neurons, at least in terms of gene regulatory responses of the neurons to dopamine receptor stimulation, is also rather uniform. However, in addition to the direct effects of dopamine on striatal output neurons, there are multiple other receptors and neuronal systems that are involved in the modulation of striatal output function. These other mechanisms produce differences in the relative responses of neurons to various inputs, including differences in the modulation mediated by the D1 and D2 dopamine receptors. The distribution of different receptors and/or their subtypes show various distributions amongst connectionally defined subpopulations of striatal neurons. In some cases the distribution patterns are similar to those of dopamine receptor subtypes, but in other cases the distribution patterns are different. An example of a receptor subtype that shows a similar pattern to the dopamine receptor distribution is the a2 adenosine receptor. The mRNA encoding this receptor 449

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has been shown to be localized specifically in neurons that also contain enkephalin mRNA, and are thus striatopallidal neurons (Ferre et al. 1993; Schiffmann and Vanderhaeghen 1993). Moreover, pharmacologic treatment with adenosine agonists has shown a specific regulation of enkephalin mRNA levels in these neurons (Ferre et al. 1993; Schiffmann and Vanderhaeghen 1993), causing similar changes in peptide as occurs through D2 dopamine receptors. Consistent with the restricted localization of this receptor to striatopallidal neurons, changes in levels of substance P, in striatonigral neurons, are not observed with the same treatments (Schiffmann and Vanderhaeghen 1993). Thus, the localization of the a2-adenosine and D2-dopamine receptor subtypes are both expressed in a similar restricted set of striatal output neurons, and activation of these receptors produce selective changes in gene regulation in these neurons. Although the changes produced by a2-adenosine and D2-dopamine receptor stimulation appear to be similar as regards changes in gene regulation of peptides in these neurons, the effects of co-stimulation of these receptors appears to be antagonistic (Ferre et al. 1993). This suggests that these two receptor systems, acting on an individual neuron, may modulate the responsiveness of these neurons to activation of the other receptor. In other cases receptors are distributed either in all striatal output neurons or in subsets of neurons that do not conform with the simple segregation of striatopallidal and striatonigral neurons. Both receptor binding studies (Herkenham et al. 1991) and in situ hybridization localization of mRNA encoding the cannabinoid receptor show that this receptor is (Mailleux and Vanderhaeghen 1992; Matsuda et al. 1993) contained in both striatal output neuron populations. Moreover, there appears to be an interaction between the activation of dopamine and cannabinoid receptors in striatal output neurons in terms of the regulation of receptor gene products (Mailleux and Vanderhaeghen 1993). Opiate receptors in the striatum have been studied for some time using receptor binding techniques (Eghbali et al. 1987; Herkenham and Pert 1982; Mansour et al. 1987; McLean et al. 1986; Tempel and Zukin 1987). Recently, the genes encoding these receptors have been identified and the coding regions sequenced, which has enabled their localization with in situ histochemistry (Evans et al. 1992; Meng et al. 1993; Thompson et al. 1993). Acetylcholine, released from interneurons within the striatum, has an important role in the regulation of striatal function. Such regulation is in part mediated through acetylcholine muscarinic receptors, which show a complex distribution pattern in striatal neuron populations. With the cloning of the family of muscarinic receptor subtypes (Bonner et al. 1987) it has been possible to localize different receptor subtypes in striatal neuron populations (Bernard et al. 1992; Weiner et al. 1990). One subtype, the m l muscarinic receptor subtype appears to be expressed by nearly all striatal medium spiny neurons. Another subtype, the m2 receptor, is expressed selectively by striatal cholinergic neurons, and may thus be an autoreceptor. Another subtype, the m4 receptor is expressed in a subpopulation that straddles the two striatal neuron populations that express D1 and D2 receptors, being contained in approximately 40% of the D2-dopamine receptor (striatopallidal) and 80% of the D l-dopamine receptor (striatonigral) neurons. Unfortunately, at this time pharmacologic agents that allow for the selective activation of the various muscarinic receptor subtypes are not available. However, it does appear that activation of these receptors has an important function in the regulation of striatal neuron activity. This may prove a complicated problem to study, as electrophysiologic studies suggest that muscarinic receptor activation may differentially alter the membrane potential of medium spiny neurons dependent on the membrane 450

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potential at the time of activation (Akins et al. 1990). Gene regulation studies also show that muscarinic agonist and antagonist treatments lead to induction of immediate early genes in subpopulations of striatal output neurons (Bernard et al. 1993). GABA and glutamate receptors are two classes of neurotransmitter receptors that are critically important to striatal function. Adequate description of these systems within the basal ganglia warrants a review that is beyond the scope of this chapter. Recent description of the distribution of the genes encoding the different subunits of both GABA and glutamate receptors are listed for reference. The GABAa receptor is composed of a combination of subunits which have been cloned and characterized (Araki et al. 1992; Seeburg et al. 1990; Shivers et al. 1989; Wisden et al. 1992; Zhang et al. 1991). The differential distribution of different subunits within the striatum suggests that this receptor system plays a complex role in striatal function. Similarly the different subtypes of glutamate receptors have been cloned, characterized and mapped within the cortex and striatum (Albin et al. 1992; Dure et al. 1992; Martin et al. 1992, 1993a, 1993b, 1993c; Petralia et al. 1994). 11.5. CELLULAR INTERACTIONS WITHIN THE STRIATUM As described above, regulation of the relative activity in striatopallidal and striatonigral neurons may be effected through the direct actions of dopamine on receptor subtypes that are differentially expressed by these two output neuron populations. However, there are multiple other neurotransmitter/receptor systems that may also function to regulate the activity of these neurons. At this time the multiplicity of interactions that presumably occur during the normal functioning of the striatum have not been worked out in any detail. Some plausible cellular interactions may be suggested based on both neuroanatomical and receptor localization studies. As described above, spiny projection neurons possess axon collaterals that extend within the striatum. Most of these appear to be distributed in a domain slightly larger than the domain of the dendritic arbors of the parent neuron. However, as seen in Figure 6 and 7, the distribution of such collaterals does not appear to cover the same area as the dendrites of its parent, and in some cases the distribution is complementary. This would suggest that one neurons axon makes contact with a neighboring spiny projection neuron, for which there is morphologic evidence (Wilson and Groves 1980). The question of whether contacts between neighboring spiny projection neurons are between neurons belonging to a similar connectionally/neurochemically defined subset of neurons or between neurons of different subsets is of some interest. Based on ultrastructural studies of the localization of peptides in boutons presynaptic to medium spiny neurons it might be suggested at least in a very preliminary way, that contacts occur between neurons belonging to the same and to different subpopulations (Bolam and Izzo 1988). Further study of this will be critical to understanding the functional significance of these local collaterals within the striatum. The neurotransmitter(s) used in these connections is also of significant interest. GABA is a likely candidate since all spiny projection neurons not only use this neurotransmitter but also possess GABA receptors. Whether the peptides that are co-localized with GABA in these neurons are employed as neurotransmitters in the connections between medium spiny neurons remains an open question. While substance P has been shown to be contained in boutons presynaptic to medium spiny neurons and to striatal cholinergic neurons, the receptor for substance P has only been localized, at this date, in cholinergic neurons (Gerfen 1991). This suggests that a single neuron might have different effects on neurons with which it makes 451

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synaptic contact dependent on the receptors on the post-synaptic neuron. In this case it is suggested that medium spiny neurons might effect cholinergic neurons through substance P-mediated mechanisms (Arenas et al. 1991), and other medium spiny neurons through other neurotransmitters, possibly GABA. The domains of the axon collaterals of medium spiny neurons are of interest in how populations of medium spiny neurons might be connected together. Most medium spiny neurons are thought to possess axon collaterals that spread within a domain roughly 200-300/zm in diamter. However, as described by Kawaguchi et al (1990), there is a subset of medium spiny neurons which have axon collaterals that spread over a considerably larger domain, up to 2 mm in diameter (see Figure 7). Such a subset of neurons has important implications for understanding the domains of populations of striatal neurons that might be functionally linked together. These neurons which have been found with such extensive local collaterals have been found to belong to the subset of striatopallidal neurons. Other characteristics of these neurons will be of great interest. Striatal interneurons undoubtedly have a major influence on the regulation of striatal medium spiny neurons, based on their synaptic contacts onto these neurons. Whether regulation of the activity in interneurons is distributed to connectionally/neurochemically defined subsets of medium spiny neurons, or are distributed more homogeneously is of interest. Most likely many combinations of interactions occur. Rather than list all the possibilities one will be suggested for which there is some experimental evidence. As described, boutons containing substance P, presumably from axon collaterals of striatonigral medium spiny neurons, make synaptic contact with cholinergic neurons, which possess the receptor for substance P (Gerfen 1991). Studies have reported substance P-mediated increase in acetylcholine release (Arenas et al. 1991) supporting the functional relevance of the neuroanatomical connections described. In addition, it has been reported that D1 receptor agonist treatment results in acetylcholine release that is mediated by substance P-receptor mediated mechanisms. Together these studies suggest that one possible cellular basis of the interaction between striatonigral and striatopallidal neurons might be mediated via connections of the striatonigral neurons with striatal cholinergic interneurons, provided that acetylcholine effects striatopallidal neurons. The select effect that stimulation of D 1 receptors has on gene regulation in striatonigral neurons and conversely that stimulation of D2 receptors has on striatopallidal neurons occurs in animal models in which dopamine is depleted from the striatum and these receptors may be stimulated independently. Of course, in the normal striatum, these receptors are most likely activated concurrently. While some models of striatal function have suggested that the interactions that occur when D 1 and D2 receptors are co-activated result from receptors co-expressed by single striatal neurons, an alternative model is that such interactions occur by way of interactions between neurons, which express predominantly one or the other dopamine receptor subtype. We have suggested some possible intercellular connections that might be involved. Moreover, these receptors are being activated in concert with other neurotransmitter/receptors expressed by striatal output neurons. Thus the effect that stimulation of any one receptor subtype, such as one of the dopamine receptor subtypes, may depend on the state of the neuron in terms of other inputs, such as glutamate inputs from the cortex, or muscarinic cholinergic inputs from striatal interneurons.

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11.6. FUNCTIONAL SIGNIFICANCE A model of basal ganglia function has been proposed based on a synthesis of experimental data which suggests that normal behavior is dependent on a balance in the output of the direct striatonigral system and the indirect, striatopallidal system (Albin et al. 1989; DeLong 1990; Mitchell et al. 1989). As described above, this model is supported by the differential effects that dopamine receptor subtype stimulation has on gene regulation in these two output systems (Gerfen et al. 1990). The model of neurologic dysfunction of the basal ganglia suggests that increased output of the indirect pathway (striatopallidal), relative to that in the direct pathway (striatonigral), results in akinesia as occurs in Parkinson's disease. Conversely, increased output of the striatonigral pathway, relative to the striatopallidal pathway, is thought to result in hyperkinetic syndromes, such as occurs in dystonia, Huntington's chorea and Tourette's syndrome, each of which is characterized by uncontrolled movement. In the case of the model of Parkinson's disease, the idea that there is an increase in the function of the striatopallidal pathway emerged from 2-deoxyglucose studies in both primates and rats which had lesions of the nigrostriatal pathway to deplete dopamine in the striatum (Mitchell et al. 1989; Trugman and Wooten 1987). In this condition, there was seen to be an increase in the glucose utilization in basal ganglia nuclei that are targets of the subthalamic nucleus, that is in the two segments of the globus pallidus (in primates) and in the substantia nigra. This suggested that increased inhibition of the globus pallidus (external segment in primates) resulting from increased striatopallidal output resulted in a disinhibition of the subthalamic nucleus and consequently increased excitatory input to the output neurons of the basal ganglia. As the output of the basal ganglia provides inhibition to the thalamus, and other targets such as the superior colliculus and pedunculopontine nucleus, increased inhibitory output was suggested to be the cause of the slowed or absent movements typical in primate models of Parkinson's disease. A test of this hypothesis was carried out by DeLong and co-workers (Bergman et al. 1990). In monkeys that showed profound bradykinesia resulting from MPTP lesions of the nigrostriatal pathway they performed lesions of the subthalamic nucleus, with the intent to block the hypothesized abnormally high output of this nucleus. Results of these lesions were dramatic in that they resulted in an immediate reversal of the lesion-induced bradykinesia. Thus, at least as far as the bradykinesia of Parkinson's disease there has been a good correlation between the connectional and neurochemical organization of striatal output pathways and dysfunctional motor behavior. As might be predicted, a simple model of basal ganglia function in the control of movement (and behavior) in terms of increased or decreased output of the striatopallidal and striatonigral pathways does not provide a full explanation of the activity in different parts of the basal ganglia during the performance of normal behaviors. On the one hand, studies of eye movements are generally referred in support of the model. Studies by Hikosaka and Wurtz (Hikosaka and Wurtz 1983a, 1983b), demonstrated that disruption of the tonic activity of the substantia nigra pars reticulata and the resultant disinhibition of neurons in the superior colliculus were tightly coupled to eye movements. However, studies of the relationship between activity in the other output nucleus of the basal ganglia, the internal segment of the globus pallidus, and movements of limb or axial musculature, do not provide such direct correlation (Hikosaka and Wurtz 1983a, 1983b; Mink and Thach 1991a, 1991b, 1991c). In some studies, most neurons recorded from the internal segment of the globus pallidus during movements show increased activity. Changes in the patterns of activity, not simple increases or decreases in the 453

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regional organization of dopamine response dynorphin

cocaine-induced c-fos 9 :i. ::5

"

)",:.i:i

: .ii: I;- .

9 :..

repeated cocaine treatment 9 . . . .

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~ :)i ~;: ~:)i~ ~~~; :~: ~:::~: : ........

Fig. 35. Diagram of the regional response within the striatum to the indirect dopamine agonist cocaine demonstrating the functional role of dynorphin in modulating this response. The basal level of dynorphin expression shows a higher level in ventral and medial striatal regions. A single injection of cocaine induces the immediate early gene c-fos by a D 1 mediated mechanism in the dorsal lateral striatal region, complementary to the area showing high levels of dynorphin. Repeated treatment with cocaine ( single daily injections of 30 mg/kg for 3 days) results in an increase in dynorphin levels in the dorsal striatal region, which has low basal expression, and a marked reduction of c-fos induction in this area, in which c-fos had previously been induced. These data suggest that dynorphin blunts the response of neurons to D 1 receptor stimulation. Further studies have shown that this effect of dynorphin is mediated through kappa opiate receptors. From Steiner and Gerfen (1993). 454

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output of the basal ganglia output are commonly reported (Filion and Tremblay 1991). In other studies, there appears to be both increased activity in the pallido-thalamic neurons, and increased activity in the target neurons of these neurons, despite the fact that stimulation of these same pallido-thalamic neurons results in inhibition in the thalamus (Anderson and Turner 1991). This finding suggests that the major target of the basal ganglia outputs in the thalamus is under the influence of other inputs that may over-ride those from the basal ganglia, possibly from the cortex. The purpose of introducing such studies is to point out that extrapolating from models of the abnormal basal ganglia to the normal function of the system requires some caution. Nonetheless, the fact that the direct and indirect striatal output pathways may, in certain conditions, be rather uniformly regulated by dopamine receptor mediated mechanisms, provides at least some insight into the functional organization of the striatum. 11.7. REGIONAL DIFFERENCES A common feature of the organization of the striatum is distinct regional and local variations in the relative expression of different neurochemical markers. The fact that these regional variations in relative expression occur in defined neuron populations that are homogeneously distributed in the striatum sets up a dichotomy that is important for understanding the functional organization of the striatum. On the one hand there are features of striatal organization that are common to all striatal regions. On the other hand, differences in the ongoing activity of inputs to different striatal regions result in differences in the level of expression of various neurochemicals that most likely reflect differential activation of the mechanisms that regulate their expression. Thus, it is important to distinguish between differences in the relative level of expression of neurochemical markers that reflect the level of activation of regulation of those markers, from the underlying mechanisms of regulation. An example of this dichotomy is the distribution of peptide markers in striatal spiny projection neurons. Striatopallidal neurons expressing enkephalin, and striatonigral neurons expressing dynorphin and substance P are fairly uniformly intermingled and evenly distributed across all regions of the striatum. This is most clearly seen using in situ hybridization histochemistry (Gerfen and Young 1988). However, even with in situ hybridization it is apparent that the relative levels of expression of the mRNAs encoding these peptides vary in different regions and in the patch-matrix compartments (discussed below). The uneveness of peptide levels is readily apparent with immunohistochemical methods. For example, the immunohistochemical localization of the peptides substance P and enkephalin are enriched in the striatal patch compartment in the dorsal parts of the striatum, and in the ventral striatum show either a more homogeneous distribution or even enriched labeling in the matrix compartment (Graybiel et al. 1981). Thus, in the normal striatum, there are distinct regional differences in the basal levels of peptide expression in striatal projection neurons. In studies in the dopamine depleted striatum, dopamine agonist treatments alter peptide levels rather uniformly in all striatal regions, independent of the regional heterogeneity that marks the normal striatum (Gerfen et al. 1990). This suggests that normal regional heterogeneity of peptide expression may reflect differences in the patterns of ongoing afferent input that regulate peptide expression, and that these patterns differ in different regions. Studies employing the indirect dopamine agonist cocaine to effect changes in striatal gene regulation reveal a possible mechanism responsible for the heterogeneity of striatal peptide expression (Steiner and Gerfen 1993). Basal dynorphin levels are relatively high 455

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in the ventral striatum, including the nucleus accumbens, and in the dorsal striatal patches. A single high dose of cocaine (30 mg/kg) results in the induction of immediate early genes, such as c-fos, in striatonigral neurons via a D1 receptor-mediated process in the dorsal striatal region that is complementary to that in which dynorphin levels are relatively high. Repeated cocaine treatment results in an elevation of dynorphin in striatonigral neurons concomitant with a decreased induction of c-fos mRNA in these neurons. This observation led to the suggestion that dynorphin functions to suppress the response of striatonigral neurons to D1 receptor activation. Consistent with this hypothesis is the finding that agonists which bind to the kappa opiate receptor through which dynorphin acts block cocaine induced c-fos mRNA. These results suggest that dynorphin may function to suppress the response of striatal neurons to the effects of dopamine mediated through D1 receptor subtype. This may be a generalized function of peptides expressed by striatal neurons, namely to provide mechanisms that modulate the responsiveness of these neurons to other neurotransmitter inputs. Differences in the normal ongoing pattern of afferent activity to the striatum may be reflected in the compensatory responses of neurons to that input. Heterogeneity in peptide levels may be the reflection of such differences. Within the striatum, there appear to be distinct differences in the dorsal and ventral patterns of expression of different peptides. This may reflect differences in either the cortical or dopamine input to these two regions. It might seem most plausible that such differences reflect differences in the dopamine innervation systems, because they are, at least in part, responsible for regulation of the peptides in question. However, for several reasons we favor as a more likely candidate the cortical or other inputs to the striatum as being responsible for such regional heterogeneity. The dopamine input to these regions originates from a continuous group of neurons in the midbrain and there is no obvious transition zone in the neurons projecting to these striatal regions. On the other hand, there are distinct differences in the cortical areas projecting to the ventral and dorsal striatum. Those projecting to the ventral striatum are for the most part allocortical or peri-allocortical areas, and include also the amygdala. Those cortical areas projecting to the dorsal striatum are neocortical areas. There are a number of major differences in the organization of these different cortical areas, such as their laminar organization, the organization of their GABA interneuron systems, and the organization of their intrinsic cortical connections, which are probably responsible for differences in the pattern of their efferent output. Whereas many of the elements of the organization of cortical-basal ganglia circuits are common in all parts of the system, other elements show distinct regional variations. In particular, there are distinct differences in the relative distribution of neurons containing the calcium binding protein parvalbumin, that show consistent regional variations in the cortex, striatum, and substantia nigra. As described in an earlier section, parvalbumin is contained in a subset of striatal interneurons, the GABAergic medium aspiny class. These neurons are most abundant in dorsolateral striatal regions (Gerfen et al., 1985), areas that receive inputs from neocortical areas. In the substantia nigra, a subset of GABA neurons in pars reticulata contain parvalbumin neurons. These parvalbumin neurons are most abundant in the regions of the pars reticulata that receive inputs from the regions of the striatum that are enriched in parvalbumin neurons. In the cortex, parvalbumin is also contained in a particular subset of GABAergic interneurons. These neurons have been well characterized by Kawaguchi and Kubota (1993) in terms of electrophysiologic properties. The number and distribution of these neurons again varies in different cortical areas, with neocortical areas showing the greatest numbers in both deep and superficial layers. These cortical areas provide inputs to the striatal 456

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regions in which parvalbumin interneurons are most abundant (Gerfen et al. 1985). Conversely, peri- and allocortical areas show a marked paucity of these neurons, as do striatal regions that receive inputs from these cortical areas, as well as regions of the substantia nigra that receive inputs from these striatal regions (Gerfen et al. 1985). Thus, it might be suggested that differences in the pattern of cortical efferent activity that is transmitted through dorsal and ventral cortico-basal ganglia circuits is related to the relative abundance of parvalbumin interneurons at each level of the system.

12. ACKNOWLEDGEMENTS

We wish to thank and give credit to H. Kita for providing examples of intracellularly filled striatal and globus pallidal neurons and to J. Tepper for similarly providing substantia nigra neurons that were used for illustrations in this paper.

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CHAPTER III

The olfactory system MICHAEL T. SHIPLEY, JOHN H. MCLEAN, LEE A. ZIMMER AND MATTHEW ENNIS

1. INTRODUCTION The olfactory system is important for reproductive/maternal functions, neuroendocrine regulation, emotional responses, aggression and for the recognition of conspecifics, predators and prey. Olfaction also plays a critical role in food selection, as the perception of flavors results from the integration of olfactory and gustatory signals. The prepotency of olfactory stimuli in memory and the control of animal behavior has long been recognized, yet the neural mechanisms that underlie these phenomena are poorly understood. Basic to the investigation of these and other olfactory functions is an understanding of the neuroanatomical and neurochemical organization of the olfactory system and its linkages to other parts of the brain. Before embarking on a systematic account of olfactory neuroanatomy and physiology, an overview of its major circuitry is presented. Odor molecules are transduced by olfactory receptor neurons (ORNs), first order neurons located in the olfactory epithelium within the nasal cavity. ORN axons project in the olfactory nerve to synaptically terminate in the olfactory bulb. The olfactory bulb contains output neurons- mitral and tufted cells - which convey olfactory information to higher order olfactory structures and to other brain systems. The relay from the nose to the mitral and tufted cells is strongly regulated by local intrabulbar circuitry and by centrifugal inputs to the olfactory bulb from other parts of the brain. Higher order olfactory structures targeted by the mitral and tufted cells include, from rostral to caudal, the olfactory peduncle (anterior olfactory nucleus), piriform cortex, olfactory tubercle, entorhinal cortex and some amygdaloid nuclei. From these primary olfactory cortical structures, further connections are made to brain regions that integrate olfactory information with other neural functions. This sketch of olfactory circuitry appears relatively simple, thus it might seem that our understanding of the functional organization of olfactory circuitry is comparable to that for other sensory systems. This is not the case, however. Indeed, there are a number of critical gaps in our knowledge of olfaction that have prevented the kinds of integrative analyses of structure and function that have led to progress in the visual, somatic sensory and auditory systems. Foremost among these gaps is our almost total ignorance of the nature of the 'olfactory code', i.e., the 'dimensions' of olfactory stimuli that are extracted and processed by the olfactory system. We do not know, for example, if there are finite 'classes' of odors comparable to primary colors. We do not know if the olfactory system extracts olfactory 'features' analogous to the orientation and/or

Handbook of Chemical Neuroanatomy, Vo112: Integrated Systems of the CNS, Part III L.W. Swanson, A. Bj6rklund and T. H6kfelt, editors 9 1996 Elsevier Science B.V. All rights reserved.

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M.T. Shipley, J.H. McLean, L.A. Zimmer and M. Ennis

velocity of visual stimuli or if the system is specialized to analyze submodalities analogous to pressure, vibration or nociception as in the somatic sensory system. How many odor selective receptor sites are on individual primary olfactory neurons and how selective are different receptors for different odors? Are some olfactory receptor neurons concerned with general properties of odors (analogous to luminance for rods in the retina) while other receptor neurons are more specialized for classes of odors, like cones are selective for wavelength? Are the olfactory receptor neurons arranged in some orderly and specific way in the olfactory epithelium and what is the relation between the location of primary neurons and their terminal projections to the bulb, i.e., is there a chemotopic or topographic organization of projections from the epithelium to the olfactory bulb? 1.1. THE OLFACTORY EPITHELIUM The sense of smell is mediated through the stimulation of olfactory receptor neurons (ORNs) by volatile chemicals. ORNs are contained in a neuroepithelium, which is located at the top of the nasal vault, along the upper portion of the nasal septum, the cribriform plate region, and the medial wall of the superior turbinate. Afferent information from these receptors is carried to the olfactory bulbs by the olfactory nerve, the first cranial nerve. In order to stimulate the olfactory receptors, airborne molecules must enter the nasal cavity, where they are subject to relatively turbulent air currents. The duration, volume, and velocity of a sniff are important determinants of an odor's stimulating effectiveness. Although these parameters differ markedly among individuals, they are quite constant for any one person. Once airborne volatiles reach the olfactory epithelium, they must pass through the layer of mucus that covers the olfactory epithelium. The stimulating effectiveness of an odor is thus also determined by the relative partitioning of the odor between air and mucus. Macromolecules in the mucus may function to bind odorants and present them to receptors; similarly macromolecules may be required to remove odorants from the receptors and or chemically inactivate odorants. The ORNs lie in a pseudostratified columnar epithelium, which is thicker than the surrounding respiratory epithelium of the nasal cavity (Fig. 1A). This epithelium rests on a vascular lamina propria. Within the epithelium are the bipolar ORNs, supporting cells (sustentacular cells), microvillar cells, and basal cells; Bowman's glands lie within the underlying lamina propria. Olfactory receptor neurons are true neurons. Their cell bodies lie in the basal two thirds of the epithelium; their apical dendrites extend to the surface. At its peripheral tip, the dendrite swells slightly to form the olfactory knob, from which several cilia extend into the mucous layer. Although human cilia do not appear to be motile, in some vertebrate species ciliary length and motility have been related to receptor age and development. Much evidence suggests that the cilia are the sites of chemosensory transduction. Basal to the ORN cell body, a nonmyelinated axon arises and joins a small bundle of other ORN axons. These axons penetrate the basal lamina, at which point the bundles become ensheathed by Schwann cells. These bundles join others to make up the 15 to 20 fascicles (fila olfactoria) of the olfactory nerve, which pass through the cribriform plate to synapse in the olfactory bulb. The supporting cells of the olfactory epithelium separate and partially wrap the ORNs. Their apical surface, in humans and some other vertebrates, is covered with microvilli, which project along with the olfactory cilia into the mucous layer. A third cell type, the microvillar cells, present at about one-tenth the number of the ORNs in 470

The olfactory system

Ch. III

A. OLFACTORY EPITHELIUM

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Fig. 1. The olfactory epithelium. A. Schematic illustration of the olfactory epithelium showing the major cell types. Inset shows the location of putative 7TM odorant receptors on cilia of ORNs. B. Hypothesized olfactory receptor-transduction mechanisms. Current evidence suggests that odor molecules bind to specific 7 transmembrane receptor (7TMr) proteins located in the cilia of ORNs. These 7 TMrs are thought to be coupled to G-proteins that activate either adenyl cyclase (AC) to generate cyclic AMP (cAMP) or phospholipase C (PLC) to generate phosphatidyl inositol (IP3). These second messengers open channels that admit calcium (Ca++) or sodium (Na+) into the cilium. These ions lead to membrane depolarization and may modulate intracellular free Ca++ levels, both of which lead to the generation of action potentials that are conducted along ORN axons to the olfactory bulb. 471

Ch. III

M.T. Shipley, J.H. McLean, L.A. Zimmer and M. Enn&

humans, have microvilli on their apical surface, projecting into the mucous layer. Their basal end tapers into a cytoplasmic extension that appears to enter the lamina propria. It is not known if this projection is an axon, although the ultrastructural appearance of the microvillar cells appears to be neuronal. Deep to the ORNs, sustentacular, and microvillar cells are the basal cells, which sit on a basement membrane just above the lamina propria. The basal cells are stem cells for the replacement of the ORNs, which in the mouse have a life span of approximately 40 days. Within the lamina propria are the secretory Bowman's glands, which provide a serous component to the mucous layer covering the olfactory epithelium. Olfactory receptor cells and axons contain olfactory marker protein (OMP), which is unique to olfactory neurons (Fig. 3A). OMP is found in a number of mammalian species, including humans, this protein, whose function is unknown, is expressed in all mature ORNs and accounts for 1% of the total protein content of these cells. Gaps in our understanding of olfactory transduction are closing rapidly. Recent breakthroughs in the cloning and characterization of seven-transmembrane receptor (7-TMR) gene families (Buck and Axel, 1991), uniquely expressed by ORNs will doubtless answer questions about the number and molecular nature of receptors that bind odorants. Thus, it will be possible to learn if individual ORNs transduce many or only a few odorants, if ORNs are 'tuned' to specific odors or classes of similar odors, if ORNs responding to similar odors are grouped together in the olfactory epithelium and/or if their axons terminate at similar target sites in the olfactory bulb. Parallel progress has also been made in understanding the transduction events that intervene between the binding of odorants to 7-TMRs and the generation of action potentials in ORNs (Fig. 1B). Several lines of evidence indicate that 7-TMRs are linked to G proteins that, in turn, activate second messenger systems such as adenylate cyclase, cAME and possibly IP3, to modulate ion channels leading to the production of action potentials that are conducted to the olfactory bulb. Thus, there is considerable optimism among olfactory researchers that long standing mysteries surrounding the molecular and biophysical events of olfactory transduction will be resolved in the coming years. However, the solution of these fundamental problems will not bring a complete understanding of the 'olfactory code' any more than the elucidation of retinal receptor transduction events has been able to clarify the neural mechanisms that underlie visual perception. The ORN is but the firstelement in a complex neural network. The operations of central olfactory networks are also poorly understood and the anatomical organization of the olfactory system appears in some respects to be fundamentally different from the familiar topographically organized circuits of the other major sensory systems. Thus, much remains to be discovered before we will approach the kind of understanding we currently have of visual, auditory and somatosensory neural network function. A better understanding of the anatomical and physiological characteristics of central olfactory circuits will provide clues about, or impose some constraints upon, the possible ways in which candidate 'olfactory codes' might be accommodated by the actual hardware of the system. Olfactory circuit organization and development have much in common with other neural systems, especially the cerebral cortex. The relatively simple cortical organization of the phylogenetically ancient olfactory system, thus, contains important clues about the most basic principles of neocortical organization. Knowledge of how such organization develops, the neural operations that are performed by olfactory circuits and of the properties of olfactory network function that have been conserved in more recently evolved cortical structures, should provide fundamental insights 472

The olfactory system

Ch. III

about the underlying computational features that have driven the selective expansion of cortical structures in the evolution of the mammalian brain. This chapter reviews the connections, chemical anatomy and physiology of central olfactory circuitry. Although great progress has been made in understanding ORNs and the olfactory epithelium, this has been the subject of several recent reviews. Greater emphasis is placed on the olfactory bulb, because it is the initial site of neural integration in the olfactory system and because it is the most thoroughly characterized central olfactory structure. Discussion of connections focuses largely on those structures most directly related to the olfactory bulb as a detailed review of circuitry beyond the primary olfactory cortex would make this chapter unwieldy. 1.2. TWO OLFACTORY SYSTEMS In macrosmatic mammals such as the rat, two components of the olfactory system are recognized; the main and accessory olfactory systems. These two components are parallel, but are, for the most part, anatomically and functionally separate. For example, ORNs in the olfactory epithelium transduce mainly volatile odors and transmit this information to the main olfactory bulb (MOB). By contrast, ORNs located in the vomeronasal organ are exposed to non-volatile odors by the engagement of a physiologically regulated pump mechanism. Axons of vomeronasal neurons project exclusively to the accessory olfactory bulb located at the dorsocaudal limit of the MOB. The central connections of the MOB and AOB to higher order olfactory structures are parallel but essentially non-overlapping. In contrast to macrosmatic mammals, microsmatic mammals such as humans have either no identifiable VNO-AOB or the VNO-AOB is only transiently present during fetal development (Humphrey, 1940; Macchi, 1951; Kreutzer and Jafek, 1980; Nakishima et al. 1985). Some mammals are anosmic (e.g. porpoises) and lack an olfactory bulb (Breathnach, 1960; Jacobs et al. 1979). Indeed, the relative size of olfactory related structures reflects the importance of olfaction to the animal. Thus, the olfactory bulb in humans is relatively small compared to the rest of the brain while the rat, which depends heavily on olfaction for reproduction and survival, has a relatively large olfactory bulb. 1.3. H U M A N DISEASES AND THE OLFACTORY SYSTEM Olfactory receptor neurons are located in the nasal epithelium. The axons of these neurons project directly to the olfactory bulb, a part of the cerebral hemisphere. This anatomical arrangement provides a direct link between the central nervous system and the organism's external environment including the possible entry of pathogens and neurotoxic chemicals into the brain (McLean et al. 1989). Until recently, olfaction has received limited attention in pathological conditions in man. Recent interest in the human olfactory system has been brought about, in part, by neuropathological investigations that described the presence of histological lesions in olfactory-related structures in Alzheimer's disease (Reyes et al. 1987; Pearson et al. 1985; Esiri and Wilcock, 1984), a condition clinically characterized by progressive intellectual decline and behavioral abnormalities. Other clinical studies have shown that olfactory deficits occur in patients with Alzheimer's disease (Doty and Reyes, 1987; Doty et al. 1988), Parkinson's disease (Doty et al. 1988) and schizophrenia.

473

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M.T. Shipley, J.H. McLean, L.A. Zimmer and M. Ennis

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2. THE MAIN OLFACTORY BULB 2.1. LAMINAR ORGANIZATION The olfactory bulb is an allocortex that, like other cortical structures, has a characteristic laminar organization. The layers of the main olfactory bulb and their principal cell types are discussed next (Fig. 2). 2.1.1. Olfactory nerve layer The superficial-most layer of the olfactory bulb is the olfactory nerve layer (ONL) which contains axons from the primary olfactory neurons (Fig. 1) and glial cells. The glial cells are thought to be derived from both the peripheral and central nervous system. At the olfactory nerve-glomerular interface, a specialized glial cell, termed the 'ensheathing cell' (Doucette, 1989), appears to provide an incomplete wrapping that separates the glomeruli from the periglomerular region (Pinching and Powell, 1971c). Recently, the deepest third of the ONL has been found to contain astrocytes. These astrocytes contain QPRT, the degradative enzyme for quinolinic acid (QUIN), a potent agonist of the N M D A receptor. QUIN, like glutamate, may arrest growth cone motility. Thus, it has been suggested that the QPRT-positive ONL astrocytes may function to degrade QUIN, thus facilitating the growth of new axons into the bulb during normal turnover and replacement of ORNs in the olfactory epithelium (Poston et al. 1991). The neuronal elements of the ONL are the axons of ORNs en route to their terminal sites in the underlying 474

The olfactory system

Ch. III

glomerular layer. The axons of primary olfactory neurons are thin and unmyelinated (Cajal, 1911; Pinching and Powell, 1971c). Candidate transmitters in olfactory receptor neurons

The possible transmitter(s) of olfactory receptor neurons have not been identified. Two molecules are relatively specific to and abundant in ORNs, olfactory marker protein (OMP) (Margolis, 1980), and carnosine (Burd et al. 1982; Sakai et al. 1988). Although OMP is abundant in ORNs (Farbman and Margolis, 1980; Monti-Graziadei et al. 1977), there is no established physiological role for this protein nor is there any indication that OMP is released by ORNs. Carnosine, a dipeptide, is present in a variety of tissues and is particularly abundant in muscle. In nervous tissue, however, carnosine appears to be uniquely expressed in olfactory receptor neurons (Ferriero and Margolis, 1975). Carnosine has some characteristics of a neurotransmitter candidate (Burd et al. 1982; GonzalezEstrada and Freeman, 1980): it is synthesized in ORNs, rapidly transported to olfactory terminals and carnosine is released by a Ca ++ dependent mechanism. To date, however, there is no consensus for a consistent postsynaptic action of carnosine. Adenosine deaminase is transiently expressed in developing olfactory epithelial cells but the functional significance of the enzyme in ORNs is unknown (Senba et al. 1987). Recently, glutamate and carnosine were co-localized in rat olfactory nerve terminals by EM immunocytochemistry (Sassoe-Pognetto et al. 1993). This suggests that glutamate, and possibly other excitatory amino acids are transmitters in the olfactory nerve. Neurophysiological studies also indicate that glutamate may be the primary excitatory transmitter in the olfactory nerve. In a turtle olfactory bulb, DNQX, a non-NMDA receptor antagonist, blocks EPSPs evoked by electrical stimulation of the olfactory nerve (Berkowicz et al. 1994). Recent studies using a rat in vitro olfactory bulb slice preparation suggest that the transmitter in the mammalian olfactory nerve may be glutamate (Ennis et al. 1994). Single shocks applied to the olfactory nerve layer (ONL) evoked (i) 1 or 2 synaptically driven spikes at short onset latencies (7-16 msec) and (ii) a longer latency burst of spikes lasting 400-1500 msec. The broad spectrum EAA antagonist, kynurenic acid, and the kainate/AMPA receptor antagonist, DNQX, completely blocked both early excitatory and burst responses. The selective N M D A antagonist, AP5, blocked the late, but not the early excitatory response component. These results suggest that glutamate and/or aspartate may be an endogenous excitatory transmitter mediating ON ~ mitral cell synaptic transmission in the mammalian main olfactory bulb. Short latency excitation of mitral cells by the olfactory nerve appears to be mediated by non-NMDA (e.g., kainate/AMPA) receptors, while longer latency and duration burst responses may be mediated by N M D A receptors. 2.1.2. Glomerular layer Immediately deep to the olfactory nerve layer is the glomerular layer (GL). The GL is one of the most distinctive structures in the brain. The glomeruli are composed of neuropil-rich spheroid structures surrounded by a distinctive shell of small neurons and glial cells (Figs. 2 and 3). The glomeruli are generally ovoid and range from 80 to 160 ~tm in diameter. Most estimates of glomerular number are similar, around 2000-3000 glomeruli/bulb for rabbits (Allison, 1949) and mouse (Allison, 1953; Brunjes, 1983; Royet et al. 1988; White, 1972). The number of glomeruli in rats has been estimated at 3000 (Meisami and Safari, 1981). It has also been estimated that there are several million 475

Ch. III

A

M.T. Shipley, J.H. McLean, L.A. Zimmer and M. Ennis

OMP

Fig. 3. Glomeruli. Photomicrographs of the main olfactory bulb showing the glomeruli that are especially well visualized by A. immunohistochemical staining with an antibody to olfactory marker protein (OMP) or B. histochemistry using cytochrome oxidase.

olfactory receptor neurons. Thus there is convergence of several thousand ORN axons in each of the 2000-3000 glomeruli, which are the initial site of synaptic integration in the olfactory system. The glomerular core is almost entirely composed of neuropil and is surrounded by a thin shell of neuron and astrocyte cell bodies. Astrocytes in the glomerular shell have a high degree of morphological specialization. The predominant type of astrocyte (termed 'wedge-shaped') has its cell body located in the glomerular shell and sends a number of thick, branched processes into the glomerular core (Bailey and Shipley, 1993). Remarkably, the processes of these astrocytes are entirely restricted to a single glomerulus. This is consistent with the hypothesis that astrocytes play a role in the formation of glomeruli. Further, as these astrocytes appear to cordon off adjacent glomeruli, this strengthens the long standing notion that each glomerulus is a discrete functional unit. The neurons of the glomerular shell are referred to as juxtaglomerular neurons (Table 1). TABLE 1. Neuron types in the main olfactory bulb Layer

Neuron type

Sizes

Cell density

GL

periglomerular short axon external tufted middle tufted Van Gehuchten mitral cell granule cell Blanes cell horizontal cell of Cajal granule cell Blanes cell Golgi cell proliferating cells

5-8/lm 8-12/lm 10-15/lm 15-18/lm 12-17/lm 20-25 r 10-16/lm 16-23 r medium 10-16 r 16-23 r 12-22/lm small

high low moderate low low very high low low low very high low low low (adult)

EPL

MCL IPL GCL

EZ

Abbreviations: GL, glomerular layer; EPL, external plexiform layer; MCL, mitral cell layer; IPL, internal plexiform layer; GCL, granule cell layer; EZ, ependymal zone.

476

The olfactory system

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B

E

F

Fig. 4. Neuron types in the MOB. Biocytin-filled cells in the MOB. A and B. Photomicrograph (A) and reconstruction (B) of a biocytin-filled tufted cell. Note the well filled apical dendritic ramifications in the glomerulus; the arrow in A indicates the initial segment of the axon. C and E show the apical and secondary dendrites of a mitral cell. D and F show a periglomerular cell. Note the extensive dendritic arbor restricted to a single glomerulus. Calibration: bars in A and D = 100/~m, bar in C -= 30/lm. The majority of the j u x t a g l o m e r u l a r n e u r o n s can be classified as one of three types: (i) small, periglomerular cells (Fig. 4E-F), (ii) slightly larger external tufted cells (Fig. 4A-B) and (iii) short axon cells (Pinching and Powell, 1971 b). The Golgi studies o f Blanes (1898), Golgi (1875) and Cajal (1911) and the Golgi and E M studies of Pinching and Powell (1971a) provide the classical descriptions o f the distribution o f j u x t a g l o m e r u l a r 477

Ch. I I I

M.T. Shipley, J.H. McLean, L.A. Z i m m e r and M. Ennis

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Fig. 5. Basic circuitry of the main olfactory bulb. Axons of ORNs form the olfactory nerve (ON). These axons terminate in the glomeruli onto mitral (M) and tufted cells (external tufted cell, ET; middle tufted cell, MT) and onto juxtaglomerular neurons including periglomerular cells (PG), ET cells and short axon cells (SA). There are one way and reciprocal synapses between the apical dendritic branches of mitral and tufted cells and the dendrites of juxtaglomerular neurons (upper inset- glomerular synapses). The lateral dendrites of mitral and tufted cells form one way and reciprocal synapses with the apical dendrites of granule cells (lower inset - dendrodendritic synapses).

References 1. Fuller, T.A. and Price, J.L. (1988); 2. Watanabe, K. and Kawana, E. (1984); 3. Halasz, N. (1987); 4. Pfister, C., Shade, R. and Ott, T. (1989); 5. Seroogy, K.B., Brecha, N. and Gall, C. (1985); 6. Matsutani, S., Senba, E. and Tohyama, M. (1988); 7. Bonnemann, C., Holland, A. and Meyer, D.K. (1989); 8. Imaki, T., Nahon, J.L., Sawchenko, P.E. and Vale, W. (1989); 9. Bassett, J.L., Shipley, M.T. and Foote, S.L. (1992); 10. Baker, H. (1986); 11. Baker, H., Kawano, T., Margolis, F.L. and Joh, T.H. (1983); 12. Baker, H., Kawano, T., Albert, V., Joh, T.H., Reis, D.J. and Margolis, F.L. (1984); 13. Baker, H., Towle, A.C. and Margolis, F.L. (1988); 14. Davis, B.J. and Macrides, F. (1983); 15. Gall, C.M., Hendry, S.H.C., Seroogy, K.B., Jones, E.G. and Haycock, J.W. (1987); 16. Halasz, N., Johansson, O., H6kfelt, T., Ljungdahl, A. and Goldstein, M. (1981); 17. McLean, J.H. and Shipley, M.T. (1988); 18. Davis, B.J., Burd, G.D. and Macrides, F. (1982); 19. Matsutani, S., Senba, E. and Tohyama, M. (1989); 20. Mugnaini, E., Oertel, W.H. and Wouterlood, F.F. (1984); 21. Macrides, F. and Davis, B.J. (1983); 22. Ribak, C.E., Vaughn, J.E., Saito, K., Barber, R. and Roberts, E. (1977); 23. Kosaka, T., Hama, K., Nagatsu, I. and Wu, J.-Y. (1988); 24. Kosaka, T., Hataguchi, Y., Hama, K., Nagatsu, I. and Wu, J. (1985); 25. Kosaka, T., Kosaka, K., Heizmann, C.W., Nagatsu, I., Wu, J., Yanaihara, N. and Hama, K. (1987); 26. Blakely, R.D., Ory-Lavollee, L., Grzanna, R., Koller, K.J. and Coyle, J.T. (1987); 27. Ffrench-Mullen, J.M.H., Koller, K., Zaczek, R., Coyle, J.T., Hori, N. and Carpenter, D.O. (1985); 28. Scott, J.W., McDonald, J.K. and Pemberton, J.L. (1987); 29. Davis, B.J. (1991); 30. Ohm, T.G., Braak, E., Probst, A. and Weindl, A. (1988); 31. Gall, C., Seroogy, K.B. and Brecha, N. (1986); 32. Sanides Kohlrausch, C. and Wahle, P. (1990b); 33. Kream, R.M., Davis, B.J., Kawano, T., Margolis, F.L. and Macrides, F. (1984); 34. Merchenthaler, I., Csernus, V., Petrusz, P. and Mess, B. (1988); 35. Tsuruo, Y., H6kfelt, T. and Visser, T.J. (1988); 36. Sanides Kohlrausch, C. and Wahle, P. (1990a). 478

The olfactory system

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cell dendrites and axons. The dendrites of the small (5-8 pm) periglomerular cells may enter more than one glomerulus but usually are preferential to one. These dendrites rarely fill the entire glomerular core instead ramifying in a discrete subregion of a glomerulus. Periglomerular cell dendrites usually have many spine-like appendages which is the chief morphological feature that distinguishes them from small external tufted cells. The short-axon cells are somewhat larger than the periglomerular cells (8-12 prn) and are distinguished in Golgi material because their dendrites are confined largely to the periglomerular shell. The axon of periglomerular/short axon cells usually course along the periphery of up to 2-4 glomeruli (Fig. 5). The external tufted cells, which are 10-15 pm in long axis, lie in the periglomerular region and usually have a single apical

TABLE 2. Candidate transmitters of olfactory bulb neurons Transmitter/ peptide

Cell type/location

Cell size

Cell number

Species

Refs.

Aspartate/Glutamate CCK

CRF

small mitral juxtaglomerular external tufted middle tufted deep tufted deep short axon vertical cells of Cajal mitral and some tufted

90 p m 11 p m 17 p m 19 p m NR NR NR NR

rat rat rat rat rat rat rat rat, monkey

1-3 4-7 4-7 4-7 4-7 4-7 4-7 8,9

DA met-ENK

juxtaglomerular G L - PG

8-13/.zm 5-8 p m

few few many many few rare rare many mitrals many many

many hamster, guinea pig

10-17 18-19

GCL GL GCL GL -

10-16 p m 5-8 p m 8-16 p m 5-8 p m

few many many many

rat, hamster many rat

20-21 22 23,24 not 20 25 25

GABA GABA + DA

- granule cells PG - granule cells PG

G A B A + parvalbumin GABA + ENK G A B A + SP + DA NAG NADPH-diaphorase NPY

E P L - external tufted G C L - granule cell G L - PG M C L - mitral cells short axon G L - PG + short axon deep G C L - short axon

small 8-16 p m 5-8 p m NR NR 14-32 p m 14-32 p m

few rare few many few few few

Somatostatin SP

G C L - PG, short axon G C L - juxtaglomerular cells EPL external tufted cells

NR 12-17 p m

few many

G C L - PG G C L - PG EPL - middle tufted cells Van Gehuchten cells

5-8 p m < 10 p m ~ 12 p m

10-15% few few

-

TRH VIP

rat rat hamster rat rat rat, human rat, human, cat marmoset rat hamster but not mouse, rat, cat, guinea pig, rabbit rat cat rat, cat

26-27 26-29 30,31 30,31,32 6,28 10,18,33

34,35 31,36

Abbreviations: CRF, corticotropin releasing factor; ENK, enkephalin; EPL, external plexiform layer; DA, dopamine; GCL, granule cell layer; GL, glomerular layer; GABA, gamma aminobutyric acid; NAG, Nacetylaspartylglutamate; NPY, neuropeptide Y; NR, not reported; PG, periglomerular; SP, substance P; T R H , thyrotropin-releasing hormone; VIE vasoactive intestinal polypeptide.

479

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M.T. Shipley, J.H. McLean, L.A. Zimmer and M. Ennis

dendrite which arborizes within a single glomerulus, although some external tufted cells have 2-3 apical dendrites that ramify in different glomeruli. External tufted cell apical dendrites are varicose but do not have spines. In addition to their thick apical dendrite(s), external tufted cells also have thinner secondary, or lateral, dendrites that extend in the external plexiform layer immediately subjacent to the glomerular layer (Fig. 4A-B). The dendritic organization of external tufted cells, thus is similar to that of the mitral cells (discussed below) although there is more variation in the number of apical and lateral dendrites of tufted than mitral cells. Some of the external tufted cells have axons that project to nearby glomeruli; these axons apparently do not enter glomeruli but, rather, terminate between glomeruli. Other external tufted cells have axons that project into the internal plexiform layer (IPL) where they travel to terminate in the IPL on the opposite side of the same olfactory bulb (Schoenfeld et al. 1985; Liu and Shipley, 1994). These 'extraglomerular' projections of the external tufted cells are discussed later. There are several other neuronal components within the glomeruli. These include dendrites of deeper tufted and mitral cells and axons from central centrifugal sources (detailed later). The studies of Pinching and Powell (1971 c) determined that ORN axons synaptically contact juxtaglomerular cells. Olfactory axons also synapse densely upon the dendrites of mitral and tufted cells. The dendrites of mitral/tufted cells and periglomerular cells also have other synaptic relationships within the glomeruli. Reconstructions of serial sections revealed that the mitral/tufted dendrites make reciprocal synaptic contacts with the dendrites and gemmules (spine-like processes) of periglomerular cells (Fig. 5; inset). These specialized reciprocal synapses are often closely associated with each other (Pinching and Powell, 1971c). Our understanding of synaptic organization in the glomeruli is far from complete because in the past decade several neurotransmitters/neuromodulators were identified in different classes of juxtaglomerular neurons. This means that the synaptic organization of glomeruli is even more complicated than originally portrayed by EM studies of the glomeruli because neurons of different transmitter phenotypes may differentially synapse with mitral/tufted cells, olfactory axons

9

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Fig. 6. Dopamine neurons in the MOB. Bright (A) and darkfield (B) photomicrographs of juxtaglomerular cells expressing the immunoreactivity to an TOH antibody. In B, there is a marked decrease in cellular expression of the dopamine phenotype due to lesion of the olfactory nerve input to the bulb using Z n S O 4. Bar in A, 100/lm.

480

The olfactory system

Ch. III

Fig. 7. GABAergic neurons in the MOB. Immunocytochemicalstaining of GABAergic cells (A) or GADcontaining terminals (B) in MOB. Bar in B, 100/lm.

and/or other periglomerular neurons. Next we review the different types of cells in the glomerular layer based on their neurochemical make-up as inferred from immunocytochemical, histochemical and in situ hybridization studies. Candidate transmitters in the glomerular layer (Table 2)

Many juxtaglomerular cells are dopaminergic (Halasz et al. 1981; Davis and Macrides, 1983; McLean and Shipley, 1988) (Fig. 6) or GABAergic (Ribak et al. 1977) (Fig. 7). In the hamster, about 70% of the DA neurons are reported to co-localize GABA while about 45% of GABAergic cells contain dopamine (Kosaka et al. 1985). In the rat, there also appears to be immunocytochemical co-localization of GABA and tyrosine hydroxylase in the juxtaglomerular cells (Gall et al. 1987; Kosaka et al. 1985). Almost all substance P immunoreactive juxtaglomerular neurons in MOB of hamster have been reported to contain both GABA and dopamine based on the presence of immunocytochemical markers for these transmitters in the same cells (Kosaka et al. 1988) although juxtaglomerular neurons in the rat do not contain substance P. Thus, some juxtaglomerular cells may contain both a catecholamine (dopamine), amino acid inhibitory transmitter (GABA) and probably also an excitatory neuropeptide (substance P), although there is considerable species variability. A few juxtaglomerular cells contain vasoactive intestinal polypeptide (Gall et al. 1986; Sanides Kohlrausch and Wahle, 1990a). Some periglomerular cells with short axons that project to the deeper granule cell layer contain NADPH-diaphorase, neuropeptide-Y (NPY) and somatostatin, which Scott et al. (1987) suggested may provide a direct route for periglomerular cells to influence granule cells. A recent study by Davis (1991) concluded that NADPH in the glomerular layer is primarily, if not exclusively contained in periglomerular cells. The 481

Ch. III

M.T. Shipley, J.H. McLean, L.A. Zimmer and M. Ennis

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The olfactory system

Ch. III

Fig. 8. CCK in the MOB. A-C. Silver-intensified CCK-immunohistochemical staining in the main olfactory bulb. CCK-immunoreactive neurons are located mainly in the superficial one-third of the EPL; the majority of these CCK-positive neurons are tufted cells. The apical and secondary dendrites of these cells are well delineated in the higher-power micrographs of (B) and (C). Thin axon-like processes course toward the IPL. In addition to the staining of cell bodies and dendrites, there is a dense, uniform CCK-like immunoreactive band consisting of terminal-lie puncta restricted to the IPL. Calibration: bar in A = 500 r bar in B = 100 /~m, and bar in C = 60/lm. (

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Fig. 9. Calcium binding proteins in the MOB. A. Distribution of cells labeled with an antibody to Calbindin (CALB) using immunocytochemistry. B. Parvalbumin (PARV) immunocytochemically labeled cells are found mainly in the EPL in the bulb. C. Golgi stained section showing impregnated external tufted cells. D. An example of how cells may be identified by other methods. In this case an external tufted ceil is shown after intracellular injection of the marker biocytin. Bar in D,/lm.

483

Ch. III

484

M.T. Shipley, J.H. McLean, L.A. Zimmer and M. Enn&

Fig. 10. Associational system in the MOB. Biocytin anterograde labeling of the intrabulbar association system (IAS) shown in dorsal (A) to ventral (F) horizontal sections. A. The injection site (asterisk) is located in the superficial half of the EPL on the lateral side of the main olfactory bulb. B-F, Biocytin-labeled tufted cells are located in the superficial part of the EPL and the deep part of the GCL. Axons and collaterals (open arrows) are densely labeled in the IPL of the medial side of the olfactory bulb ventral to the injection site. Bar in F = 800 ,um and applied to all panels.

9 + 2

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M.T. Shipley, J.H. McLean, L.A. Zimmer and M. Ennis

NADPH periglomerular cells are distinct from cells that express calbindin immunoreactivity (Alonso et al. 1993). Also reportedly present in some juxtaglomerular cells are cholecystokinin (Seroogy et al. 1985; Matsutani et al. 1988) (Fig. 8), aspartic acid (Fuller and Price, 1988; Watanabe and Kawana, 1984; Halasz, 1987), glutamate (Liu et al. 1989), thyrotropin-releasing hormone (Merchenthaler et al. 1988; Tsuruo et al. 1988) and protein kinase C (Saito et al. 1988). Calcium binding proteins have been identified in the olfactory bulb and some have been identified using immunocytochemistry in cells around the glomeruli (Fig. 9). Included in these observations are periglomerular cells which express calbindin (Baimbridge and Miller, 1982; Brinon et al. 1992; Alonso et al. 1993) and calretinin (Jacobowitz and Winsky, 1991) and superficial shortaxons cells whose dendrites do not enter glomeruli (Brinon et al. 1992). In addition, a subpopulation of small juxtaglomerular neurons are positive for acetylcholinesterase; these cells may be cholinoceptive (Nickell and Shipley, 1988). At the present time, therefore, several subclasses of juxtaglomerular neurons are distinguishable on the basis of their expression of neurotransmitters/peptides. Although these neurochemical markers serve to identify potentially different cell types, little is known of the functional significance of these transmitters/peptides for olfactory signal processing. Because most juxtaglomerular cells are very small (5-12 ~tm) and because most of them have local circuit connections, their physiological characteristics have been difficult to study.

2.1.3. External plexiform layer (EPL) Immediately deep to the glomeruli is a layer with a relatively low cell density but a very dense neuropil, the external plexiform layer (EPL). Golgi-stained sections reveal that the predominant neural elements in this layer are the dendrites of mitral/tufted and granule cells. The principal neuron types in EPL are external, middle (Fig. 12B) and deep tufted cells, named according to their relative depth in EPL, and the Van Gehuchten cells. Candidate transmitters of tufted cells and Van Gehuchten cells

There is a gradual increase in tufted cell size from the superficial to the deep parts of the EPL (Pinching and Powell, 1971b; Switzer et al. 1985). Cajal (1911) originally described the tufted cells as displaced mitral cells since they generally had the appearance of mitral cells (shaped like a Bishop's mitre). The dendritic morphologies of tufted cells vary but usually they have at least one dendrite that enters and ramifies within a glomerulus as mitral cells do. However, unlike mitral cells, many tufted cells have 2-3 apical dendrites that may enter different glomeruli. In addition, these cells have secondary dendrites that run tangentially in the EPL (Shepherd, 1972a). The secondary dendrites of tufted cells are thought to form reciprocal synapses with the apical dendrites of granule cells (Shepherd, 1972a). The middle and deep tufted cells have similar projections as the mitral cells (Schoenfeld and Macrides, 1984), thus, they may functionally be considered part of one group; the output cells of the bulb. The axons of the external tufted cells project mainly to other sites in the same olfactory bulb (Schoenfeld et al. 1985). Middle and deep tufted cells also have local collaterals in the ipsilateral bulb but most of them appear to project out of the olfactory bulb to the anterior olfactory nucleus and other rostral olfactory cortical structures (Schoenfeld et al. 1985; Scott, 1986). The intrabulbar collaterals of the superficial tufted cells form 486

The olfactory system

Ch. III

a highly organized intrabulbar network termed the intrabulbar association system (IAS). These axons project through the external plexiform and mitral cell layers into the internal plexiform layer (IPL) where they collect to form a dense tract that travels within this layer to the opposite side of the same bulb where they terminate as a dense terminal field in the IPL (Fig. 10). These tufted cells thus send a discrete, topographically organized projection to the opposite side of the same bulb. The IAS has the highest degree of point-to-point topographical organization of any known circuit in the olfactory system. C C K immunoreactive cells are mainly intrinsic to the olfactory bulb (Seroogy et al. 1985; Bonnemann et al. 1989). Recently, we have shown that the IAS is formed exclusively by CCK-containing tufted cells. Moreover, we have determined that the terminals of this CCK-ergic IAS terminate preferentially, if not exclusively, onto the apical dendrites of the granule cells (Liu and Shipley, 1994). C C K causes membrane depolarization in all neurons studied to date. Thus, it is likely that when CCK-ergic tufted cells are active, they cause depolarization of granule cells on the opposite side of the bulb. This could either increase or decrease the release of GABA depending on whether the depolarization invades the dendritic release sites or acts as a shunt for currents that would normally cause GABA release. Given the highly topographic organization of the IAS, this may lead to either highly focal inhibition or excitation on the opposite side of the bulb. A second class of EPL neurons are the Van Gehuchten cells. These cells are characterized by two or more thick primary dendrites that remain in the EPL. Axons from these cells terminate around mitral and tufted cells. Many of these Van Gehuchten cells stain positively for vasoactive intestinal polypeptide (Sanides Kohlrausch and Wahle, 1990b) calcium binding protein (Brinon et al. 1992; Alonso et al. 1993) and parvalbumin (Celio, 1990) (Fig. 9B). Table 2 summarizes transmitters/peptides observed in tufted cells. As noted, large populations of tufted cells contain the neuropeptide cholecystokinin (CCK) (Seroogy et al. 1985). One study using in situ hybridization detected substance P transcripts in some external tufted cells and in up to half of the mitral cells in MOB of rat (Warden

Fig. 11. VIP and CRF in the MOB. Photomicrographs through MOB showing cells labeled with an antibody to vasoactive polypeptide (VIP) in A and corticotrophin releasing factor (CRF) in B. The cells in A are probably Van Gehuchten cells while those in B are mitral cells. Bar in B, 100 r

487

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M.T. Shipley, J.H. McLean, L.A. Zimmer and M. Ennis

and Young, 1988), but to date no studies using immunocytochemistry have detected substance P in mitral cells in any species (Baker, 1986; Inagaki et al. 1982; Schults et al. 1984). Even in the hamster, which has many substance P juxtaglomerular cells in MOB, substance P is not present in the mitral/tufted cells, at least as detected by immunocytochemistry (Baker, 1986). Many of the middle tufted cells are reported to contain vasoactive intestinal polypeptide (VIP) in the rat (Fig. l lA) (Gall et al. 1986), but in the cat, this peptide appears to be in van Gehuchten cells not tufted cells (Sanides Kohlrausch and Wahle, 1990a). In the hamster, a few middle tufted cells are NADPH diaphorase positive (Davis, 1991).

2.1.4. Mitral cell layer (MCL) Deep to the external plexiform layer is the mitral cell layer (Fig. 2). This is a thin layer that contains the somata of mitral cells (25- 35/2m diameter) arranged in almost a monolayer. These cells are the principal output cells of the bulb and, with some minor species differences (cf. Scott, 1986), have one apical dendrite that enters a single glomerulus, where it branches extensively and is synaptically contacted by olfactory axons (Shepherd, 1972a) (Figs. 4C,E, 5 and 12). Intracellular filling of impaled neurons with HRP was used to describe the detailed structure of individual neurons (Mori et al. 1983; Kishi et al. 1984; rabbit; Mori et al. 1981 b; turtle). In mammals, each mitral cell sends a single primary dendrite to a single glomerulus. (In the turtle bulb mitral cells typically send a primary dendrite to two glomeruli; Mori et al 1981b). In addition to this primary dendrite, each mitral cell also sprouts several secondary dendrites. These secondary dendrites extend for very long distances through the EPL, often covering 2 mm or 1/4 of the bulb's circumference. Reconstruction of mitral cells did not reveal any preferential orientation of the dendrites; the arbors tended to be symmetrical. Scott's laboratory studied a sample of mitral and tufted cells (Orona et al. 1983; rat) using extracellular iontophoretic deposits of HRP among the dendrites of mitral/tufted cells. Dendrites passing through the injection site took up the tracer, which was transported throughout the neuron, providing Golgi-like labeling of a small population of neurons. These workers describe two classes of mitral cells. Secondary dendrites of the most numerous class (Type I) ramify almost entirely in the deep part of the EPL. A second class of mitral cells (Type II) sends dendrites almost exclusively to more superficial parts of the EPL. The distributions of the dendrites of these two types of mitral cells correspond with the laminar distribution of cytochrome oxidase activity in the EPL (Mouradian and Scott, 1988). The superficially located dendrites of type II mitral cells extend over a smaller area than the deeper dendrites of type I mitral cells. The population studied by Orona et al. may not be a random sample, since visualization of neurons was dependent upon the placement of the iontophoretic injection; however, in their population 79 of 274 mitral cells (29%) were classed as 'Type II'. As it courses through the EPL, the apical dendrite of mitral and tufted cells receives very few synapses and the main shaft of the apical dendrite appears to be insulated by glial sheaths (Shipley and Zahm, 1990). The secondary dendrites of mitral cells engage in dendro-dendritic synapses with dendrites of granule cells. In addition, they may receive centrifugal and Van Gehuchten cell inputs (Jackowski et al. 1978; Rall et al. 1966).

488

Fig. 12. Golgi impregnated cells in the olfactory bulb. A. The large arrow indicates the soma of a mitral cell. One can follow (small arrows) the primary dendrite to a glomerulus. B. A middle tufted cell. C. The large arrow indicates a mitral cell while the small arrows show granule cells with their dendrites coursing towards the EPL. P

00

W

9 * 2

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M.T. Shipley, J.H. McLean, L.A. Zimmer and M. Ennis

Responses of mitral cells to odors

Responses of mitral cells to odors are typically complex: during odor exposure, units may be initially excited then inhibited, inhibited then excited, or may exhibit more complex responses. The character of these responses may alter with odor concentration. Individual units may more reliably discriminate between odors if unit activity is recorded in relation to an artificial sniff cycle (Macrides and Chorover, 1972). Recent authors have emphasized the necessity of testing the response of each cell over a range of odor concentrations (Meredith, 1986; Harrison and Scott, 1986). Testing with several odors, each at several different concentrations, showed that a significant number of cells respond differently to at least two odors at all odor concentrations (Wellis et al. 1989). Similar results in the salamander led to the concept of 'concentration tuning' of bulb units: individual cells appeared to respond best to a particular concentration of each odorant (Kauer, 1974). If an odor causes strong excitation in a particular region of the bulb surface, then there ought to be lateral inhibition of surrounding cells. Meredith (1986) found a few pairs of cells, separated by relatively large distances, which gave approximately opposite responses to odor stimulation. Pairs of this type appear not to be numerous, however. On the other hand, closely neighboring mitral cells, presumed to connect with the same glomerulus, showed strong correlations in their responses to odors (Buonoviso and Chaput, 1990). Candidate transmitters of mitral cells

One proposed transmitter for mitral cells is N-acetyl-aspartyl-glutamate (NAG) (Table 2) (Blakely et al. 1987; Ffrench-Mullen et al. 1985). NAG has been observed in mitral cells using immunocytochemistry (Blakely et al. 1987). However, a recent neurophysiological study cast doubt on a transmitter role for NAG (Whittemore and Koerner, 1989) in mitral cells (see below, 2.5.4. Transmitter(s) mediating MOB to PC monosynaptic excitation). A few, unusually small, mitral cells appear to contain aspartate and project to the piriform cortex (Fuller and Price, 1988). Many mitral cells, as well as tufted cells in the EPL, have been reported to express glutamate immunoreactivity (Liu et al. 1989). Recently, the neuropeptide, corticotropin releasing factor (CRF), has been demonstrated in mitral and some tufted cells using both immunocytochemistry (Fig. 11B) and in situ hybridization in the rat (Imaki et al. 1989). CRF fibers were also observed in the molecular layer of the piriform cortex. This finding is consistent with CRF being a releasable neural peptide in mitral cells since mitral cells synaptically terminate in the molecular layer of the piriform cortex. A similar localization of CRF has been reported in the squirrel monkey suggesting that this peptide may be a conserved transmitter/ modulator in the mitral/tufted cells of many mammals (Bassett et al. 1992). Finally, calretinin, a calcium binding protein, has been shown by immunohistochemistry to be localized in mitral cells (Jacobowitz and Winsky, 1991). Transmitter candidates for mitral and tufted cells are discussed further in 2.5.3., Projections to olfactory cortex.

2.1.5. Internal plexiform layer (IPL) Immediately subjacent to the mitral cell layer is the internal plexiform layer (IPL) (Fig. 2). The IPL is a thin layer with a low density of cells but with many axons and dendrites. Golgi studies indicate that the IPL contains the axons of mitral/tufted cells, dendrites 490

The olfactory system

Ch. III

of granule cells and axons from other unidentified sources. Some axons in IPL originate from raphe nuclei (5-HT) (McLean and Shipley, 1987b), locus coeruleus (NE) (McLean et al. 1989) and the nuclei of the diagonal band (ACh) (Shipley et al. 1986) (see Section 5). As noted, the IPL also contains a rich plexus of CCK containing axons and terminals (Liu and Shipley, 1994) which derive from CCK containing tufted cells. The IPL also contains a population of multipolar neurons, larger than granule cells, that express ACHE. As these neurons do not express CHAT, the synthetic enzyme for ACh, but lie in a layer richly targeted by centrifugal inputs from the nucleus of the diagonal band, we have suggested that theses cells may be cholinoceptive.

2.1.6. Granule cell layer (GCL) The granule cell layer is the deepest neuronal layer in the bulb (Fig. 2). This layer contains many small (8-10/2m in long axis) granule cell neurons. Frequently, 3-5 granule cells are arranged in row-like aggregates of tightly packed somata. Granule cells in these aggregates are coupled by gap junctions, which may serve to synchronize the functional activity of these neurons (Reyher et al. 1991). Granule cells have also been found mixed with mitral cells in the mitral cell layer. Golgi studies indicate that granule cells lack axons. They have basal dendrites that ramify in the GCL and a thicker and longer apical dendrite that enters and ramifies extensively in the EPL. Freeman (1972a) noted that the dendrites of Golgi-impregnated granule cells tend to be of the same length; thus, deeply situated granule cells will tend to interact with mitral dendrites deep in the EPL, while superficially situated granule cells will interact principally with mitral and tufted cells having dendrites in the superficial parts of the EPL. This laminar segregation of the granule cell connections was studied more rigorously by Orona et al. (1983), again using the technique of localized extracellular injections of HRP. According to their reconstructions, each granule cell sends one or two dendrites toward deeper parts of the GCL, about 50-100 r and a single apical dendrite extends superficially toward the EPL. This dendrite generally does not branch until reaching the EPL; it may not branch extensively until it reaches the superficial EPL. The dendritic arbor of the granule cell extends over an area of diameter 100-200 r However, not all granule cells follow this pattern. The exceptions include granule cells whose dendrites project deeper, toward the center of the bulb and granule cells with dendrites that do not reach the external plexiform layer (Schneider and Macrides, 1978). In addition, there are also non-granule cells in the GCL, including the Blanes cells. The laminar distribution of granule cells and their apical dendrites, combined with the segregation of mitral and tufted cell secondary dendrites into different depths of the EPL (Orona et al. 1984) implies that mitral and tufted cells interact with different populations of granule cells. Tufted cells projecting from the bulb terminate primarily in the anterior olfactory nucleus (AON) and anterior piriform cortex (Scott et al. 1980; Scott, 1981; Schneider and Scott, 1983). Reciprocal projections from the AON, anterior piriform cortex and the anterior commissure terminate primarily in the IPL and superficial GCL and thus primarily influence superficial granule cells whose dendrites most probably contact tufted cells. Mitral cells project to more caudal levels of piriform cortex and the reciprocal projections from posterior piriform cortex terminate more deeply in the GCL (Davis and Macrides, 1981; Luskin and Price, 1983b). Thus, Orona et al. (1983) propose that there may be two parallel and partially independent olfactory processing systems in the bulb: the mitral cells and their associated granule cells, and the tufted cells, with their separate population of granule cells. It is not clear, however, to what extent 491

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M.T. Shipley, J.H. McLean, L.A. Zimmer and M. Ennis

the segregation of granule cell contacts is common to all mammals. In the rabbit, Mori et al. (1983) describe a class of granule cells with short dendrites; these cells have their cell bodies near the MCL and project into the deep parts of the EPL. Granule cells appear to be electrotonically short (Rall and Shepherd, 1968; Shepherd and Brayton, 1979); thus, depolarization of any part of the cell spreads relatively effectively to all parts. Most granule cells appear not to be spontaneously active in the absence of olfactory or electrical stimulation, as passage of a recording electrode through the GCL ordinarily encounters no active units; however, spontaneously active units which may be granule cells can be isolated near the MCL (McLennan, 1971). However, granule cells can produce spikes during excitation by antidromic (LOT), or orthodromic (anterior commissure) stimulation (Mori and Kishi, 1982). In a population of 20 granule cells studied with intracellular recording, 8 responded to odor stimulation with spikes, 7 responded with excitatory responses, 4 showed no response and 1 cell was inhibited (Wellis and Scott, 1990). Candidate transmitters of the granule cell layer

There are extensive synapses between the dendrites of the granule cells and the secondary dendrites of mitral/tufted cells (Fig. 5; inset). Mitral/tufted cell dendrites make excitatory synapses onto the dendrites of granule cells and the dendrites of the granule cells make inhibitory synapses onto the dendrites of mitral/tufted cells (Shepherd, 1972). Most of the granule cells contain GABA (Ribak et al. 1977) (Fig. 7) which inhibits mitral and tufted cells (Jahr and Nicoll, 1982; Shepherd, 1972a,b). Some peptides have also been localized in other cells in the GCL. For example, a few cells containing NPY are found in this layer in the rat (Gall et al. 1986) and human (Ohm et al. 1988). In the rat, the axons of NPY cells appear to ramify in the more superficial layers such as the glomerular layer (Gall et al. 1986). These neurons are probably short-axon cells as opposed to granule cells. Granule cells are both intensely and lightly stained with an antibody to calretinin (Jacobowitz and Winsky, 1991). 2.1.7. Mitral-granule cell interactions: Anatomical considerations

Some calculations based on the dimensions of mitral and granule cell dendrites provides insight into their possible functions. Each Type I mitral cell sustained an average of 6.2 secondary dendrites, each of which branches several times (Orona et al. 1983). There are approximately 25 mitral cells projecting into a single glomerulus (Allison and Warwick, 1949). If the secondary dendrites of these mitral cells extended radially outward equally in all directions there would be 150 secondary dendrites from each glomerular unit, providing 1 secondary dendrite per 2.4 degrees. At 2 mm from the center of the glomerular unit, secondary dendrites from one glomerulus would be separated by about 80 r Since each secondary dendrite branches several times, dendrites may be even more closely spaced. A similar conclusion can be reached from the estimate of Mori et al. (1983) of the mean total length of the secondary dendrites of a single mitral cell as 15,000 r 25 mitral cells would have a total dendritic length of 375,000 r (375 mm). If these dendrites were cut into 2 mm lengths and distributed evenly over a 2 mm square the average spacing would be 11 r Since the diameter of the entire dendritic arbor of a granule cell is 100-200 r (Orona et al. 1983), each granule cell could be contacted by dendrites of several mitral cells from each glomerulus. Thus, if the mitral cells of a single glomerulus are activated or inhibited as a unit and the secondary 492

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dendrites of these mitral cells sustain action potentials, their activation could create a uniform excitation of granule cells over a large part of the bulb. 2.1.8. Subependymal zone

The deepest layer in the main olfactory bulb is the subependymal zone, a cell-poor region in the adult. The cells in this layer line the ventricle (if present) and during development the progenitors of many olfactory bulb cells derive from this zone, although other olfactory bulb neurons (granule cells) may be generated in more caudal forebrain regions then migrate into the bulb in what has been termed 'the rostral migratory stream' (Altman, 1969; Kishi, 1987; Shimada, 1966). 2.2. TRANSMITTER RECEPTORS IN THE MOB (Table 3) 2.2.1. Excitatory amino acids (EAAs)

Based on pharmacological studies, four major groups of EAA receptors are reported in the CNS: kainate, NMDA, AMPA, and metabotropic receptors. All four receptor subtypes have been identified in MOB, and the laminar specificity of these receptors have been reported. Kainate receptors are restricted to the MCL, EPL and IPL in the bulb (Gall et al. 1990; Miller et al. 1990; Monaghan and Cotman, 1982; Wisdon and Seeburg, 1993). NMDA receptors are found in every region of MOB except for the IPL (Monyer et al. 1994; Petralia et al. 1994; Watanabe et al. 1993). Receptors of the AMPA subtype are localized in the MCL, IPL, and glomerular layers (Molnar et al. 1993; van den Pol et al. 1994; Martin et al. 1993; Petralia and Wenthold, 1992). Immunocytochemical evidence suggests that these receptors are localized to the processes and cell bodies of periglomerular, mitral, and tufted cells (Molnar et al. 1993; Petralia and Wenthold, 1992). Subunits for the metabotropic glutamate receptor (mGluR) are localized in the GL, MCL, and GCL (Ohishi et al. 1993; Martin et al. 1992; Shigemoto et al. 1992; Shigemoto et al. 1993; Tanabe et al. 1992; Takaaki et al. 1992). Immunocytochemical studies have localized mGluR's to periglomerular and mitral cell bodies, while mGluR mRNA has been identified in tufted cells (Martin et al. 1992; Shigemoto et al. 1992). 2.2.2. GABA receptors receptors are present in every layer of MOB except for the IPL (Fritshy et al. 1989; Laurie et al. 1992; Palacios et al. 1981; Zhang et al. 1991; Persohn et al. 1992; Bowery et al. 1987; Richards et al. 1987; Chu et al. 1990). While mRNA studies localize GABAA subunits to mitral, tufted, and periglomerular cells, immunocytochemical studies report only mitral and granule cells to contain GABAA (Fritshy et al. 1989; Laurie et al. 1992). GABAB receptors are restricted to the glomerular layer in MOB (Chu et al. 1990; Bowery et al. 1987). Modulatory transmitter receptors in MOB are discussed in Section 5. GABA A

2.3. INFLUENCE OF THE OLFACTORY NERVE ON TRANSMITTER EXPRESSION IN MOB NEURONS Input from the olfactory epithelium is important for the normal expression of transmitters in many types of olfactory bulb neurons. For example, the olfactory nerve influences 493

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M.T. Shipley, J.H. McLean, L.A. Zimmer and M. Ennis

the expression of the dopamine (DA) phenotype in juxtaglomerular neurons. If the olfactory epithelium is destroyed by detergent o r ZnSO4, or if functional activity in the pathway is perturbed by closure of the nares, then tyrosine hydroxylase (TH) immunoreactivity is lost in dopaminergic juxtaglomerular neurons (Fig. 6). This phenomenon of transneuronal regulation of transmitter phenotype has been demonstrated in adult rats (Kawano and Margolis, 1982; Baker et al. 1983; Baker et al. 1984), mice (Nadi et al. 1981; Baker et al. 1983), dogs (Nadi et al. 1981), hamsters (Kream et al. 1984) and developing rats (McLean and Shipley, 1988). Thus, these cells appear to require the

TABLE 3. Transmitter receptors in the M O B Receptor mAChR1 mAChR2 mAChR3 mAChR4 nAChR alphal alpha2 beta 1 beta2

GL

+ +

+ +

EPL + + + + +

MCL

+ + + + +

IPL + + + +

+ + + +

GCL

References 3,5,8,27,28,32

++

+ +

1,18,29,36,40

++

++ ++

++

++

++

NR

+ + NR

Ol

D2

+ + (ONL) + + (GL)

5-HTIA 5-HT2AJr

NR

Kainate NMDA AMPA Metabotropic

+ + + + + +

GABAA GABAB

+ + + +

+ + + +

+ + + +

+ + + + + +

NR + + + + + +

+ +

+ + NR

+ +

+ +

7,11,12,14,15, 16,17,19,22, 23,30,31,33, 34,35,37,38

+ + NR

2,4,6,9,20,21, 26,41,

+ +

NR

10,13,20,21,39

Key: + +, receptors present; - - , receptors absent; NR, not reported Abbreviations: MOB, main olfactory bulb; GL, glomerular layer; ONL, olfactory nerve layer; EPL, external plexiform layer; MCL, mitral cell layer; IPL, internal plexiform layer; GCL, granule cell layer; mAChR, muscarinic cholinergic receptor; nAChR, nicotinic cholinergic receptor; alpha and beta adrenergic receptors; D, dopamine receptors; 5-HTIA and 5-HT2A/C, serotonin receptors; N M D A , N-methyl-d-aspartate receptor; AMPA, ~-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid receptor; GABAA/B, gamma aminobutyric acid receptor. References 1. Booze et al., 1989; 2. Bowery et al., 1987; 3. Buckley et al., 1988; 4. Chu et al., 1990; 5. Fonseca et al., 1991; 6. Fritschy et al., 1989; 7. Gall et al., 1990; 8. Hunt and Schmidt, 1978; 9. Laurie et al., 1992; 10. Liu and Shipley, in preparation; 11. Martin et al., 1992; 12. Martin et al., 1993; 13. McLean et al., 1994; 14. Miller et al., 1990; 15. Molnar et al., 1993; 16. Monaghan and Cotman, 1982; 17. Monyer et al., 1994; 18. Nicholas et al., 1993; 19. Ohishi et al., 1993; 20. Palacios et al., 1981; 21. Persohn et al., 1992; 22. Petralia and Wenthold, 1992; 23. Petralia et al., 1994; 24. Pompeiano et al., 1992; 25. Pompeiano et al., 1994; 26. Richards et al., 1987; 27. Rotter et al., 1979; 28. Sahin et al., 1992; 29. Sargent-Jones et al., 1985; 30. Shigemoto et al., 1992; 31. Shigemoto et al., 1993; 32. Spencer et al., 1986; 33. Takaaki et al., 1992; 34. Tanabe et al., 1992; 35. van den Pol et al., 1994; Wanaka et al., 1989; 37. Watanabe et al., 1993; 38. Wisden and Seeburg, 1993; 39. Whitaker-Azmitia et al., 1993; 40. Young and Kuhar, 1980a; 41. Zhang et al., 1991.

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epithelial input in order to maintain TH and, hence, express their DA phenotype. The loss of TH immunoreactivity is not due to cell death because the neurons can be detected with antibodies to other DA enzymes (Baker et al. 1984). Moreover, Baker (1990) recently showed that GABA continues to be expressed in juxtaglomerular neurons in which TH is downregulated following deafferentation. Epithelial input is also required in order to trigger the initial developmental expression of TH (McLean and Shipley, 1988; Baker and Farbman, 1993). Thus, both the developmental induction and maintenance of the DA phenotype depends on the presence and normal function of the olfactory nerve. It is not known if the influence of olfactory epithelial input on the DA neurons is by direct synaptic contact or through less direct means such as transneuronal regulation of mitral/tufted cell dendrites which could then influence DA expression in the PG cells (McLean and Shipley, 1988). As mentioned above, Pinching and Powell (1971 a) showed that there is direct contact of PON axons with PG cells, however, it is not known if olfactory axons synapse on DA juxtaglomerular cells. Recently it was demonstrated that the predominant DA receptor in the bulb is of the D 2 subtype. Surprisingly, this receptor is localized not only in the glomerular layer, as expected, but equally densely in the olfactory nerve layer (Nickell et al. 1990). When the olfactory epithelium is destroyed by ZnSO4, all D2 receptor binding is eliminated in both the olfactory nerve layer and in the glomeruli (Shipley et al. 1991). This raises the interesting possibility that the DA neurons are presynaptic to olfactory nerve terminals. More direct evidence for this hypothesis comes from the recent demonstration that olfactory neurons contain abundant levels of D 2 receptor mRNA transcripts (Shipley et al. 1991). Thus, while it has been presumed for two decades that olfactory nerve terminals do not receive presynaptic contacts, it is possible that juxtaglomerular dendrites release DA which acts on olfactory terminals but without a classic synaptic specialization [see below, section 2.4.3]. In this scenario, the loss of TH in juxtaglomerular DA neurons following deafferentation might reflect a down regulation of DA synthesis due to the loss of the postsynaptic targets (ORN terminals) of the DA neurons. While deafferentation does not influence the expression of GABA in periglomerular neurons, other transmitters are influenced by the olfactory nerve. In the hamster, many juxtaglomerular neurons express substance P and this peptide is downregulated following deafferentation (Kream et al. 1984). In addition, new results indicate that the influence of the olfactory nerve also extends to the expression of CCK in tufted cells and to CRF in mitral cells (Shipley et al. unpublished observations). The influence of the olfactory nerve on olfactory bulb cells has been studied in other ways. Laing and colleagues (Laing et al. 1985) found that exposing animals to pure air decreased the size of mitral cells, while exposure to specific odors over prolonged periods caused regional increases in mitral cell size. These findings were interpreted to suggest that odor quality may be coded at the level of the mitral cells (Panhuber and Laing, 1987). Light and dark granule cells have been described (Struble and Walters, 1982) and these two subpopulations differ in their reaction to unilateral nares occlusion (Skeen et al. 1985; Frazier and Brunjes, 1988)" the lighter granule cells found in the deeper regions of GCL are the first to be affected by nares closure. Frazier and Brunjes (1988) speculate that the lighter cells are affected first because they arrive first in GCL. Nares occlusion reduces the number of both light and dark granule cells and it has been speculated that the reduction is due to cell death rather than decreased cell proliferation (Kaplan et al. 1985; Frazier and Brunjes, 1988) since the granule cells continue to be born in the proliferative subependymal zone (Frazier and Brunjes, 1988). 495

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2.4. FUNCTIONAL ORGANIZATION OF THE MOB 2.4.1. Organization of olfactory nerve inputs to MOB Analysis of the functions of the olfactory bulb would be greatly simplified if the coding of information in primary olfactory axons were known. Unfortunately, widely differing concepts of olfactory coding can be supported by available information. 2.4.2. Broad topographic mapping A mapping of odors might be created either by segregation of receptor subtypes sensitive to classes of odors into different regions of the epithelium (Kauer and Moulton, 1974; Mackay-Sim et al. 1982) or by physical mechanisms which segregate odorants according to their physical properties onto regions of the epithelium (Mozell, 1964, 1966, 1970; Moulton, 1976) or some combination of both. Either of these mechanisms could generate spatial and temporal patterns which might be conveyed to the olfactory bulb by topographic ordering of olfactory nerve axons. However, topographic order in the projection from the olfactory epithelium to the bulb is crude relative to that of other sensory systems (e.g. Land and Shepherd, 1974; Land, 1973; Astic and Saucier, 1986; Saucier and Astic, 1986; Costanzo and O'Connell, 1980; MacKay-Sim and Nathan, 1984; Dubois-Dauphin, 1981; Duncan et al. 1990). This modest topographic ordering might, however, be sufficient to map a small number of epithelial regions onto the bulb surface; topographic ordering of odor properties might be sharpened by reorganization of olfactory axons within the olfactory nerve before termination in the bulb (Kauer, 1987) or by bulb circuitry. Recently, Buck and Axel (1991) have shown that members of the multi-gene family encoding putative 7-TM olfactory receptors are selectively expressed in 3-4 anatomically distinct zones in the olfactory epithelium. These 'expression zones' are roughly organized as rostrocaudal strips. Thus, there may be at least a crude topographical organization of receptor gene sub-families in the olfactory epithelium. However, the interpretation of these findings is uncertain as it is still unclear whether distinct 7-TM receptor gene products recognize specific odors or, alternatively, if different receptors recognize different sites on odorant molecules. In other words, is there one receptor for each odor or are many different receptors required to distinguish among different odors? Glomeruli as functional units

The lack of point to point topography in the projection of the olfactory epithelium to the olfactory bulb and the early recognition that olfactory bulb glomeruli appear to represent anatomically discrete units has led to the widespread view that glomeruli play an important role in olfactory coding. The precise relationship of olfactory receptor axons to individual glomeruli has, thus, drawn considerable attention. Axons from the olfactory epithelium travel in parallel fascicles (Daston et al. 1990) but form a dense plexus after they pass through the cribriform plate. Individual glomeruli receive fascicles entering from different directions (Le Gros Clark, 1951; 1956). A similar plexus of fibers is seen in amphibians (Scalia, 1976; Kauer, 1981; Duncan et al. 1990). These early observations of Le Gros Clark were confirmed by later studies using degeneration or tracer methods to label the fibers arising from restricted regions of the

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epithelium (Land and Shepherd, 1974; Land, 1973; Adamek et al. 1986; Stewart and Pedersen, 1987). Within the broad regions of the bulb labeled after focal injections in the epithelium, a few densely labeled glomeruli are generally surrounded by many lightly labeled glomeruli (Adamek et al. 1986). Within the olfactory nerve layer the divergence of olfactory axons is limited: A small cut across the olfactory nerve layer results in a relatively narrow band of denervation caudal to the cut (Wellis and Scott, 1989); mitral cells beneath the denervated region are not responsive to nerve stimulation rostral to the cut. Freeman (1972c, 1974d) reached a similar conclusion by mapping the extent of field potentials and of excited mitral cells following focal electrical stimulation of the nerve layer. The analysis of this anatomical arrangement by Le Gros Clark (1951; 1956) cannot be much improved: 'One possibility is that there are different types of olfactory receptors which are differentially susceptible to various forms of olfactory stimulus, and that impulses from the several categories of receptor become segregated and conveyed to different glomeruli of corresponding categories .... Another possible interpretation of the resorting of olfactory nerve fibres is that those derived from equivalent regions of the complicated turbinal system.., are regrouped so as to be led to different sets of glomeruli. ... However, there is another possible explanation of the olfactory nerve plexus on the surface of the bulb - that it serves precisely the opposite purpose of ensuring a complete randomization of fibres so that each glomerulus receives impulses from every type of receptor; in other words, that each glomerulus is in itself a functional unit concerned with the total range of olfactory discrimination .... In any case, whatever the peripheral analytical mechanism for olfactory discrimination may be, on anatomical grounds the glomeruli of the olfactory bulb appear to provide a suitable basis as a series of central analyzers, for.., each glomerulus with its connecting mitral cells forms a virtually closed system' (Le Gros Clark, 1956). Thus, the anatomy of the olfactory nerve suggests two models of olfactory coding in which the glomeruli are important functional units. The first assumes that each glomerulus receives fibers specific for some particular odor 'quality'. One implication of this model is that the amount of the olfactory bulb required to represent all odors would be proportional to the number of distinct olfactory receptor types present in the olfactory epithelium. If there are a relatively small number of olfactory receptor types, a relatively small part of the bulb would contain a neural representation of all possible odors. This can be illustrated by a simple calculation. There are about 2000 glomeruli in the rabbit olfactory bulb (Allison and Warwick, 1949). If there were 7 classes of receptors, and if glomeruli receive axons from only one class of receptors, then each class of receptor would project to about 300 different glomeruli. If these glomeruli were scattered randomly over the bulb surface, and the average glomerulus were 200 r in diameter, then on average a region of the bulb surface of about 0.2 m m 2 (0.5 mm diameter) would contain one of each type of glomerulus. However, larger numbers of receptor types would require a proportionally larger part of the bulb surface to represent all possible odors. For a very large number of odor qualities, the entire bulb would be required to discriminate between odors. In the second model suggested by Le Gros Clark, each glomerulus receives a sample of many different receptor types. Random distribution of individual fibers would result in a broad representation of different fiber types within glomeruli. Each glomerulus would respond to many possible odors. In this case small portions of the bulb would be capable of representing all possible odors. Experiments by Slotnick et al. (1987) have shown that a learned olfactory discrimina497

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tion survives ablation of most of the bulb. This supports the idea that a relatively small part of the bulb is capable of representing a wide range of odor qualities. On the other hand, several kinds of experimental evidence indicate that glomeruli are heterogeneous functional units. Metabolic activity in individual glomeruli, determined by 2-deoxyglucose (Lancet et al. 1982) or by cytochrome oxidase activity (Fig. 3) (Shipley and Costanzo, 1984; Costanzo et al. 1984; Weinberg and Meisami, 1989; Mouradian and Scott, 1988), is constant within a glomerulus but varies with no apparent pattern from glomerulus to glomerulus. Also consistent with the importance of glomeruli as functional units is the observation that a slow electrical potential recorded across a single glomerulus showed a different pattern of response to a battery of 9 odorants for each site tested (Leveteau and MacLeod, 1966). Finally, an important consideration for any model of olfactory coding is the fact that primary olfactory neurons undergo turnover and replacement throughout life (Moulton, 1974; Graziadei and Metcalf, 1971). Unless the replacement axons re-establish similar functional connections, perceived odor perception would be altered and the animal would lose the ability to remember odors. As this is not the case, it is clear that the olfactory pathway does have the ability to re-establish appropriate functional connections. This might be accomplished by the maintained expression of biochemical markers which guide new axons of a particular type to their appropriate termination. An alternate hypothesis is suggested by the observation that regenerating olfactory neurons exhibit increased odor selectivity after establishing contact with the olfactory bulb (Mair et al. 1982). This suggests that primary olfactory neurons might have relatively unspecified odor selectivity until their terminals reach the glomerular layer of the olfactory bulb; selectivity could be conferred by molecular signals retrogradely transferred from target cells in the bulb to olfactory receptor neurons in the epithelium.

2.4.3. Neural processing in the glomerular layer Summation of inputs The firing rate of a primary olfactory neuron is presumably proportional to the concentration in its vicinity of the odorants to which it is sensitive. Even for an ideal transducer, however, there is a necessary minimum of noise arising from the random variation of odorant concentration in a small region (Schild, 1988). There are, however, almost certainly a large number of olfactory neurons associated with each class of odorant and the sum of the activity of all or part of these neurons would be a far more precise indicator of odorant concentration. Thus, one plausible function of the initial synaptic connection in the bulb is the derivation of an average activity of input fibers. If there are 50,000,000 olfactory receptors and 2000 glomeruli (Allison and Warwick, 1949), then each glomerulus receives 25,000 olfactory axons. If there are 25 mitral cells associated with each glomerulus (Allison and Warwick, 1949), then the average number of axons terminating on each mitral cell is about 1000. Electrotonic conduction within the apical dendrite of a single mitral cell provides an adequate mechanism for summation of the inputs to that neuron. The reduction in noise in the summated activity is 21000 or about 30-fold (Holley and Doving, 1977; Schild, 1988). If a glomerulus receives axons of a single odorant class, noise in the summated activity of these axons would be reduced by the 225,000 or 160 fold from the noise present in a single axon. It is not clear, however, how much of this enormous potential increase in sensitivity is realized as there is no evidence that the mitral cells associated with a single glomerulus project to common 498

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targets (Scott et al. 1980; Scott, 1981; Ojima et al. 1984). Hence, there appears to be no opportunity for additional convergence of the mitral cells from a single glomerulus onto a relatively small number of target neurons, as would appear necessary for effective summation of their activity.

Local circuit organization in the glomerular layer In addition to summation of inputs from olfactory nerve terminals, a second function of the circuitry of the glomerular layer is regulation of transmission from the olfactory nerve to mitral and tufted cells. The anatomy of the glomerular layer, reviewed above, provides a basis for such regulation (Fig. 5). Terminals of the olfactory nerve synapse both with the apical dendrites of mitral and tufted cells and with juxtaglomerular (JG) cells. JG cells, which send processes primarily into a single glomerulus, form reciprocal synapses with the apical dendrites of mitral and tufted cells; many of these are symmetrical and, therefore, are considered inhibitory. There is thus a 'feedforward' pathway for inhibition of mitral/tufted cells: olfactory nerve terminals excite JG cells which, in turn inhibit mitral/tufted cells. The mitral/tufted side of the reciprocal synapses are asymmetrical and are, therefore, considered excitatory. Thus there is also a pathway for 'feedback' inhibition from mitral and tufted cells back to JG cells. Some JG neurons send axons to neighboring glomeruli but it is not known if these local projections are excitatory or inhibitory (Schneider and Macrides, 1978).

Neurophysiology of JG cells Extracellularly recorded units in the glomerular layer fire one or a few spikes following electrical stimulation of the olfactory nerve (Getchell and Shepherd, 1972b; Freeman, 1974b). Intracellular recordings from a small population of PG neurons showed that ON stimulation produces an EPSP of about 30 msec duration (Wellis and Scott, 1990). In some cells, larger EPSPs resulted in fewer spikes, presumably because of depolarization block. In extracellular recordings this depolarization block would be interpreted as inhibition. JG neurons are facilitated following weak shocks to the olfactory nerve; at longer stimulus intervals or with stronger stimuli, these neurons are inhibited (Getchell and Shepherd, 1975b). Facilitation and inhibition were inferred from changes in response latency, response threshold, or the number of spikes generated by a constant stimulus. Intracellular recordings from JG cells, revealed an IPSP of 200 msec duration produced by ON stimulation (Wellis and Scott, 1990). There is only limited information on the intracellular responses of JG cells to odors (Wellis and Scott, 1990); of three cells, all responded to odor stimulation with a burst of spikes associated with a depolarization. Two of the three showed a reduction in response with successive 'sniffs', although there was no apparent IPSP.

Glomerular inhibition Olfactory nerve stimulation activates both glomerular (JG and mitral/tufted cells which are reciprocally connected) and infraglomerular (Mitral/granule/mitral) inhibitory systems. Therefore, it is difficult to separate the relative contribution of these two inhibitory systems to mitral cell inhibition. However, stimulation of the olfactory nerve (orthodromic stimulation) results in a longer lasting inhibition of mitral cells than stimulation 499

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of LOT (antidromic stimulation) (Freeman 1974a; Nickell et al. 1994). Conditioning shocks given to ON inhibit responses to test shocks; this inhibition has an exponential time course of hundreds of msec, substantially longer than the duration of spike activity generated in PG neurons by ON stimulation (Freeman, 1974b; Nickell et al. 1994). This inhibition was not activated by antidromic (LOT) stimulation (Freeman, 1974a: Nickell et al. 1994), indicating that it is not associated with the mitral/granule inhibitory system which is activated by antidromic stimulation. Thus, activation of interneurons in the glomerular layer may cause a relatively slow inhibitory process which reduces transmission from olfactory nerve terminals to mitral/tufted cells. Transmitter mechanisms of glomerular inhibition

Additional clues about the functions of glomerular circuitry are provided by the localization of transmitters and receptor subtypes within the bulb: PG cells contain the transmitter GABA (Ribak et al. 1977; Mugnaini et al. 1984). PG and other JG cells contain dopamine (Halasz et al. 1978,1981; Davis and Macrides, 1983; McLean and Shipley, 1988). GABA and DA are co-localized in some JG cells (Gall et al. 1987; Kosaka et al. 1985). Receptor subtypes for these transmitters many of which use relatively slow 'second messenger' mechanisms are present in the glomerular layer. GABAB receptors are located in the glomerular layer and nowhere else in the MOB (Bowery et al. 1987). Topical application of baclofen inhibits the field potential response to ON stimulation and inhibits mitral cell responses to nerve stimulation in olfactory bulb slices in the rat (Nickell et al. 1994). This supports the idea that GABAB receptors are involved in the regulation of transmission from the olfactory nerve to mitral and tufted cells. DA receptors in the bulb are the D 2 subtype (Nickell et al. 1991) and exclusively located in the glomerular layer and the olfactory nerve layer (Nickell et al. 1991). D 2 and GABAB receptors have been shown to inhibit neurons by similar second-messenger mediated mechanisms (Pinnock, 1984; Gahwiler and Brown, 1985; Innis and Aghajanian, 1987; Lacey et al. 1988). Earlier observations suggested that there is presynaptic inhibition of olfactory nerve terminals (Freeman, 1974a; Mori et al. 1984; Jahr and Nicoll, 1981). Jahr and Nicoll (1981) monitored currents flowing into the terminals of the frog olfactory nerve. Electrical stimulation of the nerve produces a prolonged depolarization of the terminals. Jahr and Nicoll considered this effect to be analogous to the depolarization associated with presynaptic inhibition in dorsal roots of the spinal cord (review, Nicoll and Alger, 1979). Now it appears that DA presynaptically inhibits transmission from olfactory nerve terminals to mitral/tufted and possibly other JG cells. R e c e n t l y , D 2 receptor mRNA was found to be expressed in the olfactory epithelium; removal of the olfactory bulb which causes all olfactory receptor neurons to degenerate, eliminated D 2 mRNA expression from the olfactory epithelium (Shipley et al. 1991). This demonstrates that D2 receptor mRNA is localized to olfactory receptor neurons. Destruction of the olfactory epithelium eliminated all D 2 binding in the olfactory bulb (Shipley et al. 1991). This indicates that all D 2 receptors in the bulb are localized to olfactory receptor axons and terminals. Taken together, these results strongly suggest that DA released from JG cells acts o n D 2 receptors on olfactory nerve terminals. Preliminary electrophysiological experiments in vivo and in an in vitro slice preparation support this hypothesis (Zimmer et al. 1995): Either DA or D2 agonists block transmission from the olfactory nerve to mitral cells; D 2 antagonists block this action of DA.

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Excitatory systems of the glomerular layer There are several morphological cell types within the glomerular layer (Schneider and Macrides, 1978; Macrides et al. 1985); any of these might provide excitatory inputs to mitral and tufted cells, although there are no available data on the synaptic connections of these subtypes. There is also evidence for excitatory interactions within the glomerular layer. Electrical stimulation of the olfactory nerve activates some units which should be outside the spread of the activated fibers and some units are activated after long latencies suggestive of polysynaptic pathways (Getchell and Shepherd, 1975b; Wellis and Scott, 1990). In turtle bulb, orthodromic but not antidromic stimulation activates a slow excitatory current (Nowycky et al. 1981). Freeman (Freeman, 1974a,b; Martinez and Freeman, 1984) has argued that JG cells are primarily excitatory. The most direct evidence is the demonstration of a potential difference indicating current flow between the glomerular and external plexiform layers. This potential is activated by orthodromic (olfactory nerve) but not by antidromic (LOT) stimulation and is of relatively long duration (30-50 msec). It is argued that this potential is the result of excitatory input to the apical dendrites of mitral cells. In summary, the available evidence suggests that both excitatory and inhibitory actions are present within the glomerular layer; further work is needed to define the roles of these events in signal processing in the glomeruli.

2.4.4. The mitral/granule cell inhibitory system Much experimental work has focused on the reciprocal synaptic connections between the secondary dendrites of mitral/tufted cells and the granule cells because the density of these synaptic interactions suggests their fundamental importance to bulb function.

Lateral inhibition versus self-inhibition The responses of the mitral/granule system to single or paired orthodromic and antidromic volleys are well understood. But, the function of this reciprocal synaptic mechanism in processing odor evoked activity depends on its response to spatially and temporally patterned activity in the olfactory nerve. One possible function of the mitral/granule system is to mediate some form of lateral inhibition which would sharpen the differences between active and inactive regions of the bulb, a function analogous to similar operations in the retina and other sensory systems. In the classical descriptions of the mitral/granule system, however, mitral cells are described as primarily self-inhibitory (Rall et al. 1966; Rall and Shepherd, 1968; Rall, 1972; Shepherd, 1972a). Self-inhibition would predominate if there are many reciprocal synapses near the mitral soma or if the secondary dendrites are electrotonically short (so that synaptic currents occurring anywhere on the dendrite are conducted with relatively little decrement to the soma). If, on the other hand, [i] the secondary dendrites are electrotonically long, [ii] there are relatively few reciprocal synapses near the soma, and [iii] the dendrites sustain action potentials, then inhibitory currents generated in distal parts of the dendrite would have less effect on the soma, but action potentials in the soma might propagate out the dendrite, activating the reciprocal synapses with granule cells. In this case an active mitral cell (or group of cells) would preferentially inhibit other mitral cells in the more remote reaches of its dendritic arborization. To the degree that one or the other of these two modes of inhibition 501

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predominates during the processing of odor signals, very different kinds of functions could be performed by the mitral/granule system. As noted above, the secondary dendrites of mitral cells extend for extraordinary distances, on the order of 2 mm. With reasonable values of membrane resistance, the secondary dendrites would be electrotonically long; activity in the reciprocal synapses would decrease length constants further. Thus, voltage changes in distal parts of the dendrites are probably only weakly coupled to the soma. Inhibitory activity in more proximal parts of the dendrite soma would be more strongly coupled to the soma. Both self- and lateral-inhibition may be generated by this anatomical arrangement. The relative importance of the two kinds of inhibition can be assessed by intracellular recordings from mitral cells and data from such recordings are available from several studies. The inhibitory potentials (IPSPs) recorded intracellularly from mitral cells following LOT stimulation increase in a graded manner as LOT antidromic stimulation strength is increased. The appearance of a spike in the impaled cell does not cause a large increment in the IPSP (Yamamoto et al. 1962; Mori and Takagi, 1978), suggesting that most of the IPSP results from activity in other mitral cells. In the isolated turtle olfactory bulb individual mitral cells were impaled and stimulated by injection of current through the recording pipette (Jahr and Nicoll, 1982a). In nearly all cells tested stimulation of the impaled cell alone produced a smaller IPSP than either orthodromic (ON activation) or antidromic (LOT) stimulation. There was considerable variability, however: in some cells direct stimulation of the impaled neuron produced a large IPSP but in many others only a small IPSP was produced. These observations suggest that both self- and lateral-inhibition are present in the mitral/granule system. The rate of spikes from the soma of a mitral cell probably reflects the summation of excitatory currents from the primary dendrite and inhibitory currents from proximal parts of the secondary dendrites. An action potential generated in the soma probably propagates out the secondary dendrites, possibly as a calcium spike (Jahr and Nicoll, 1982a). This propagated spike will release an excitatory transmitter, but the quantity of transmitter released and the extent of spread of the spike will depend upon the local level of inhibition. The IPSP (in the dendrite) following the spike blocks further release of transmitter for a period of several msec. The activity of other mitral cells and of centrifugal inputs to the bulb (see below) may also set independent levels of inhibition in the distal parts of the secondary dendrites. Thus, quite complex temporal and spatial patterns could be generated. The effect of the mitral/granule system on inputs to the bulb can be indirectly assessed by determining the pattern of excitation and inhibition caused by localized stimulation of the olfactory nerve. Freeman (1974c) mapped the pattern of average evoked potential (AEP) and single unit responses following focal stimulation of the olfactory nerve. This study concluded that ON stimulation results in an initial excitatory focus which is replaced by a larger inhibitory focus and not in an excitatory focus surrounded by an inhibited region, as expected for pure lateral inhibition. Oscillation and the mitral/granule system

The modes of operation of the mitral/granule system are too complex to be adequately summarized as self- or lateral inhibition. An adequate description of the behavior of this system may require the development of models with appropriate simplifying assumptions. One such model is based on the ability of the mitral/granule system to develop oscillations. 502

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Oscillatory activity of about 50 Hz was a prominent feature of early descriptions of the response of the olfactory bulb to odors (Adrian, 1950). Depth recordings demonstrate that the oscillations recorded at the bulb surface are the result of synaptic currents flowing into granule cells following mitral cell activation (Freeman, 1972a). Stimulation of LOT or ON at low stimulus amplitudes results in oscillatory averaged responses rather than the monophasic responses observed after large shocks (Freeman, 1972b). Antidromic activation of mitral cells by stimulation of LOT at frequencies near this natural oscillation result in alternation of large and small responses (Mori et al. 1977; Westecker, 1970). The generation of these oscillations can be rationalized from the classical descriptions of the mitral/granule system (Shepherd, 1972a,b, 1979; Rall, 1972). An alternative viewpoint is provided by noting that the mitral/granule system constitutes a negative feedback loop (Freeman, 1972b). All such feedback systems tend to oscillate at the frequency corresponding to the total of delays around the loop; thus, the mitral/granule system may be expected to oscillate at a frequency corresponding to the duration of the IPSP plus the delay in the EPSP which generated it. The duration of the IPSP is strongly influenced by anesthetics; in unanesthetized preparations, or with certain anesthetics, this period is approximately 20 msec, corresponding to the experimentally observed oscillation frequency of 50 Hz. Barbiturate anesthesia, which increases the delay around the loop by prolongation of the IPSP (Stewart and Scott, 1976; Scott and Stewart, 1979; Nicoll, 1972b) reduces the frequency of the spontaneous oscillations (Adrian, 1950; Scott and Aaron, 1977). This oscillatory behavior provides one model for the operation of the mitral/granule system. If a spatially patterned input is applied to the bulb, groups of neighboring mitral cells receiving sufficient excitatory input might break into oscillation; interactions mediated by the long secondary dendrites would tend to synchronize these groups of neighboring cells over a large portion of the bulb surface. Mitral cells receiving insufficient excitation for oscillation would be inhibited by the activity of the oscillating cells. This mode of operation could accommodate either the broad topographic or the glomerular models of olfactory input. If olfactory information is coded on broad regions of the bulb then the mitral cells within a region receiving sufficient excitation might oscillate; the long secondary dendrites might serve to inhibit other regions receiving less excitatory input. Alternatively, if olfactory input is coded into individual glomeruli, the long secondary dendrites might function to synchronize oscillations between active but spatially separated glomeruli. 2.4.5. Glomerular versus infraglomerular inhibition

It has long been recognized that inhibition plays a key role in olfactory bulb network function. The classical mitral-granule-mitral loop which provides feedback regulation of mitral cell excitability has been the subject of much study. By, contrast, there has been less consideration of intraglomerular inhibition in models of olfactory circuit function. Intraglomerular inhibition regulates sensory throughput at the first olfactory synapse and, because it precedes and is in series with the mitral-granule-mitral system, it influences the operations of the mitral-granule-mitral loop. In light of these considerations, it may be worthwhile to reconsider the functional significance of the mitral-granulemitral system. If orthodromic transmission is regulated by an intraglomerular system with a longer time constant than the granule to mitral synapse, what might be the function of the 503

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M.T. Shipley, J.H. McLean, L.A. Zimmer and M. Enn&

mitral-granule-mitral loop? While it may function to further limit orthodromic excitability, its primary role may be in the centrifugal regulation of olfactory bulb function. Centrifugal feedback projections arise in virtually all structures targeted by mitral/tufted cells, including the anterior olfactory nucleus, and piriform, periamygdaloid and lateral entorhinal cortex. The majority of these feedback projections terminate with excitatory synapses on the granule cells. Thus, centrifugal projections to the bulb comprise a massive negative feedback system that operates through the granule-mitral circuit. Thus a primary function of the classical granule-mitral system may be to mediate centrifugal feedback inhibition while the glomerular inhibitory system functions more as an intrabulbar feedforward mechanism that regulates orthodromic activity. Regardless of the cellular localization of the intraglomerular GABAB receptors, the strength of the sensory input should regulate the amount of negative gain applied to subsequent input. The intraglomerular inhibitory system, thus, may regulate the dynamic range of the olfactory system such that mitral/tufted cells are less prone to 'saturation' by strong input from the olfactory nerve. 2.5. OUTPUTS OF THE MOB 2.5.1. Intrabulbar collaterals

The axons of the mitral cells give off collaterals within the bulb in the internal plexiform and granule cell layers (Mori et al. 1983). The main axons course predominantly in the lateral olfactory tract which forms at the level of the AOB. These caudally directed axons give off collaterals in the anterior olfactory nucleus (AON) and other regions of olfactory cortex (Figs. 13, 18, 19). Tufted cells collateralize to an even greater extent in the bulb than mitral cells. The intrabulbar association pathway formed by CCK-ergic tufted cells was discussed earlier. If the reciprocal Synapses between mitral secondary dendrites and granule cells are suppressed by a conditioning antidromiC shock to LOT, then the response to a test LOT shock is positive in the GCL (Nicoll, 1970), as would be expected from activation of mitral axon collaterals synapsing on granule cells in the GCL. These connections may thus provide a pathway for mitral cell activation of granule cells which is not affected by inhibition of mitral cell secondary dendrites. Tufted cells respond more reliably and more vigorously to odors or to electrical stimulation of the olfactory nerve than mitral cells (Schneider and Scott, 1983). 2.5.2. Mitral/tufted cell projections beyond the MOB

The primary output of MOB is through mitral cells and the middle and deep tufted cells. Some of the outputs of the MOB have a modest degree of topographical organization. For example, neurons in the dorsolateral quadrant of MOB project to the dorsal part of the external subdivision of AON while output cells of the ventral half of MOB project to the lateral subdivision (Schoenfeld and Macrides, 1984). Output neurons of MOB that project to more caudal regions (e.g. PC) are evenly distributed in MOB. Intracellular injections of HRP into mitral cells have been used to show the arborizations of individual axons. The axons often have many collateral terminal arbors in AON and fewer in the more caudally located piriform cortex (Ojima et al. 1984). The terminal arbors have a patchy anterior-posterior distribution in layer Ia of AON and piriform cortex, which according to Ojima and colleagues (1984; rabbit) is reminiscent to the 504

Ch. III

The olfactory system

OLFACTORY CORTEX

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Fig. 13. Basic olfactory network. Schematic of the networks linking the olfactory bulb and primary olfactory cortex. Olfactory nerve axons (ON) terminate in the glomeruli (glom) onto mitral (m) and tufted (t) cells which project via the lateral olfactory tract (LOT) to layer Ia of primary olfactory cortex to terminate on the dendrites of layer II-III pyramidal (p) cells. Layer II-III pyramidal cells in rostral olfactory cortex project to layer Ib in caudal olfactory cortex and vice versa. Olfactory cortical pyramidal cells also send reciprocal projections back to the olfactory bulb. Thus olfactory bulb output is continuously modified by feedback from areas it targets. Inhibitory interneurons in olfactory bulb and olfactory cortex (shown in gray) modulate network function. Neurons in the ipsilateral (AONi) and contralateral anterior olfactory nuclei (AON) link olfactory networks in the two hemispheres via the anterior commissure. patchy distribution of thalamic input to the visual cortex. Some mitral cells branch to project to both olfactory cortex and the olfactory tubercle. Mitral cells that are close together are reported to have similar patterns of axonal projections to the olfactory cortex (Buonviso et al. 1991). While there are some hints of organization of MOB output projections, the preponderance of evidence from a large number of studies using a range of tract tracing techniques indicates that the outputs of the bulb do not have the kind of point to point topographical projections to their target structures as is characteristic of other sensory systems. 2.5.3.

Projections

to olfactory

cortex

M O B projects to several structures of the ipsilateral hemisphere, including the superficial plexiform layer of the anterior olfactory nucleus, piriform, periamygdaloid and lateral entorhinal cortices, taenia tecta, the anterior hippocampal continuation, indusium griseum and the olfactory tubercle (Figs. 14, 18, 19). Collectively, the regions directly innervated by the output of the MOB have been referred to as primary olfactory cortex (De Olmos et al. 1978). Most of these projections have been reported in several species (for review, cf. Shipley and Adamek, 1984). A recent study has also shown a 505

Ch. III

M.T. Shipley, J.H. McLean, L.A. Zimmer and M. Ennis MAIN OLFACTORY SYSTEM PRIMARY OLFACTORY GORI"EX

:"ACCESSORYOLFACTORYCORTEX

ACCESSORY OLFACTORY SYSTEM Fig. 14. Major connections of the main (MOB) and accessory (AOB) bulbs with cortical (gray panels) and subcortical structures (ellipses). Output projections of MOB and AOB shown by solid lines; reciprocal and centrifugal projections to MOB and AOB are shown by gray lines. Cortical areas comprising the primary and accessory olfactory cortex are indicated by squares. Abbreviations: ACo = anterior cortical amygdaloid nucleus; AOB - accessory olfactory bulb; Me = medial amygdaloid nucleus; A O N = anterior olfactory nucleus (m = medial division); PCo = posterior cortical amygdaloid nucleus; BST = bed nucleus of the stria terminalis; D H R - dorsal hippocampal rudiment; D P C = dorsal peduncular cortex; D R - dorsal raphe nucleus; Ent = entorhinal cortex; LC = locus coeruleus; LPO = lateral preoptic area; MOB = main olfactory bulb; M n R - median raphe; BAOT = nucleus of the accessory olfactory tract; N L O T - nucleus of the lateral olfactory tract; DB = nucleus of the diagonal band; PeCo = periamygdaloid cortex; Pir = piriform cortex; Tu = olfactory tubercle; TT = taenia tecta

direct MOB projection to the supraoptic nucleus (Smithson et al. 1989). The primary efferent pathways of the main olfactory system are summarized in Figures 14 and 19. The terminal zones of bulb outputs are discussed below in the section on primary olfactory cortex.

2.5.4. Transmitter(s) mediating MOB to PC monosynaptic excitation The transmitter(s) mediating excitatory transmission from the olfactory bulb to PC is the subject of considerable interest. EM and Golgi studies indicate that MOB inputs in layer Ia synapse on the basal dendrites of layer II pyramidal neurons and on semilunar neurons (Haberly, 1983). MOB terminals make asymmetrical synaptic contacts with dendritic spines, suggesting that they are excitatory synapses. Consistent with this finding, activation of bulb inputs to PC via electrical stimulation of the olfactory bulb or by direct stimulation of LOT produces rapid monosynaptic depolarization and spiking in superficial pyramidal cells. The fast excitatory transmission at this synapse suggests that it is mediated by an EAA. Anatomically, it has been difficult to delineate 506

The olfactory system

Ch. III

an EAA in this pathway due to the problems associated with immunocytochemical detection of EAAs although a recent study suggests that mitral cells contain glutamate (Liu et al. 1989). However, neurochemical studies demonstrate that bulbectomy reduces evoked release of aspartate and N-acetylaspartylglutamate in PC (Collins and Probett, 1981a; Ffrench-Mullen et al. 1985) and stimulation of LOT induces a calcium-dependent release of glutamate and aspartate in PC (Bradford and Richards, 1976; Collins, 1979; Collins and Probett, 1981b). Receptor localization studies indicate that the three main classes of EAA receptors, as well as the metabotropic glutamate receptors are present in PC, although information about their precise laminar distribution is not available. Neurophysiological studies suggest that the initial MOB-evoked excitation of superficial pyramidal cells is mediated by several classes of EAA receptors. Such excitation is attenuated by 2-amino-4-phosphonobutyric acid (AP4) (Ffrench-Mullen et al. 1986; Hori et al. 1982; Collins, 1982; Ffrench-Mullen et al. 1985; Hasselmo and Bower, 1991), a proposed antagonist of the metabotropic glutamate receptor. AP4 has been reported to block actions of glutamate at an unidentified postsynaptic EAA receptor and is thought to decrease glutamate release via actions at a presynaptic metabotropic glutamate receptor. In agreement with this hypothesis, Hasselmo and Bower (1991) reported that AP4 decreased afferent fiber-evoked excitation of PC neurons through a presynaptic mechanism. Collins and Richards (1990) recently found that protein kinase inhibitors reduce LOT-evoked monosynaptic excitation of PC, suggesting a role for a metabotropic, second messenger-mediated component of excitatory transmission at this synapse. LOT-evoked excitation of PC is also blocked by DNQX (Collins and Buckley, 1989), a potent postsynaptic antagonist of kainate and AMPA receptors. By contrast, selective NMDA receptor antagonists do not reduce LOT-evoked monosynaptic excitation of PC cells. 2.6. CENTRIFUGAL AFFERENTS TO MOB Centrifugal afferent inputs to the olfactory bulb are very dense, come from multiple sources (Figs. 14, 18, 19) and play important roles in regulating neural processing in the olfactory bulb. These inputs display some degree of anatomical organization and neurochemical heterogeneity. The centrifugal afferents may be divided into two groups: (i) afferents from olfactory related structures and (ii) non-olfactory subcortical modulatory afferents to the olfactory bulb. These are distinguished because the olfactory related projections almost certainly mediate specific olfactory sensory and association functions whereas the modulatory afferents have widespread projections that influence CNS functions across neural system lines. Olfactory-related centrifugal afferents arise from many sources including the AON, piriform cortex, periamygdaloid cortex, entorhinal cortex, nucleus of the lateral olfactory tract and amygdala. Subcortical modulatory afferents originate in the brainstem and basal forebrain. Olfactory related centrifugal afferents to the main olfactory bulb are discussed below in Section 3.3.2. The organization of subcortical 'modulatory' inputs to the bulb are discussed below in Section 5.

3. PRIMARY OLFACTORY CORTEX

The main olfactory bulb projects to a collection of structures referred to collectively as primary olfactory cortex (De Olmos et al. 1978). These structures may be usefully divided into three groups: (A) the anterior olfactory nucleus (Fig. 15); (B) rostral olfac507

Ch. III

508

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The olfactory system

Ch. III

tory cortex comprising the indusium griseum, anterior hippocampal continuation, taenia tecta (Figs. 15 C-D, 16), infralimbic cortex (Figs. 15D, 16) and the olfactory tubercle (Fig. 17 A and B) and (C) lateral olfactory cortex (Fig. 17) comprising, from rostral to caudal; piriform, periamygdaloid, transitional and entorhinal cortices.

3.1. ANTERIOR OLFACTORY NUCLEUS (AON) 3.1.1. Architecture of AON

Caudal to the olfactory bulb is a distinctive structure, the anterior olfactory nucleus (AON). Herrick (1924), who first coined the term anterior olfactory nucleus, regarded it as a nucleus. However, the AON is a laminated structure, consisting of a plexiform layer and a rather homogeneous layer of tightly packed cells except in the center (Haberly and Price, 1978a) Golgi and Nissl studies have shown that AON is laminated and contains pyramidal cells (Haberly and Price, 1978a). Many of the output cells in the different subdivisions of AON are pyramidal, save for neurons in the external subdivision. Based on these architectonic features, most workers today consider AON to be a cortical structure. The AON has been divided into several subdivisions based on cytoarchitecture and connections (Haberly and Price, 1978a; De Olmos et al. 1978) (Fig. 15). In this chapter we do not review the cytoarchitectural criteria used to distinguish subdivisions of the AON; for interested readers these subdivisions have been described previously (De Olmos et al. 1978; Shipley and Adamek, 1984; Haberly and Price, 1978b). 3.1.2. Inputs to AON

Other than inputs from MOB, AON inputs arise from other subdivisions of the ipsilateral AON and from the contralateraI AON (De Carlos et al. 1989; Luskin and Price, 1983b). In addition, there are inputs from piriform cortex and entorhinal cortex (Luskin and Price, 1983a,b), anterior amygdaloid area (De Carlos et al. 1989; Luskin and Price, 1983a,b), posterolateral cortical amygdala nucleus (De Carlos et al. 1989; Luskin and Price, 1983b), olfactory tubercle (Luskin and Price, 1983), nucleus of the lateral olfactory tract (De Carlos et al. 1989) and the temporal portion of CA1 division of the hippocampus (van Groen and Wyss, 1990), infralimbic region of the medial prefrontal cortex (Takagishi and Chiba, 1991) and 'modulatory' inputs from the raphe nuclei (McLean and Shipley, 1987b), locus coeruleus (McLean et al. 1989) and nucleus of the diagonal band (De Carlos et al. 1989). Our knowledge of these complex patterns of connections is summarized in Figures 27 and 28 of the original publication by Haberly and Price (1978b). 3.1.3. Outputs of AON

All subdivisions of the ipsilateral AON project to both the ipsilateral and contralateral main olfactory bulbs except the external division (AON pars externa, AONpE) which projects only to the contralateral MOB (De Olmos et al. 1978) (Fig. 18A). The AON contains the largest number of neurons projecting to the bulb from any one source (Carson, 1984a). The core of the AON is white matter, analogous to the white matter of the cerebral cortex. The axons of AON neurons collect in this white matter to project either to the 509

Ch. III

M.T. Shipley, J.H. McLean, L.A. Zimmer and M. Ennis

ipsilateral bulb or to continue caudally where they join the anterior commissure to cross the midline and terminate in the contralateral AON and/or bulb. There is some degree of laminar topography of the terminal projections to MOB from different AON subdivisions, in rat (Luskin and Price, 1983b; Reyher et al. 1988) and hamster (Davis and Macrides, 1981). In both species, the ventral and posterior subdivisions of AON project bilaterally to the GCL (mainly the superficial half) and the deep third of the GL. In the hamster, the lateral and dorsal divisions of AON project mainly to the superficial GCL and GL (ipsilateral) with no projection to the glomeruli, while in the rat the dorsal division terminates evenly in the GCL and lightly in the EPL but not to the GL. The external subdivision of AON in both species terminates heavily in a thin band just deep to the IPL in the contralateral MOB. Luskin and Price (1983b) described additional outputs of the AON including projections to ventral taenia tecta, piriform cortex and olfactory tubercle, endopiriform cortex, ventral agranular insular cortex and nucleus accumbens (from ventroposterior portion of AON). According to Luskin and Price, there are very few projections outside olfactory cortex from AON, although Price et al. (1991) have confirmed a strong projection from AON to the lateral hypothalamus.

3.1.4. Organization of AON circuitry A ON pars externa, the intrabulbar associational system and topography in the olfactory system

One of the most puzzling but consistent observations about the olfactory system is the lack of point-to-point spatial topography in the connections from the epithelium to the bulb and from the bulb to higher olfactory cortical structures. This lack of topographical organization is in marked contrast to other sensory systems for which there is typically a precise point-to-point mapping of the receptor surface across several synaptically linked stages of the sensory pathway. This 'diffuse' organization has been interpreted to mean that the neural coding of 'odor space' does not depend in any straightforward way on an orderly spatial arrangement of olfactory receptor neurons in the epithelium or upon their orderly mapping into the glomerular layer of the olfactory bulb. This may be premature as the mapping of the epithelium to the bulb has been suggested by several studies to consist of both diffuse and focal components; the focal components may represent some degree of topography (Adamek et al. 1986; Land, 1973; Land and Shepherd, 1974; Stewart, 1985). Recent anatomical studies have demonstrated a previously unsuspected degree of point-to-point topography for some inter- and intrabulbar circuits. A subpopulation of external tufted cells send axons to the opposite side of the same bulb (Schoenfeld et al., 1985; Liu and Shipley, 1994). This intrabulbar association system (IAS) is discretely organized; neighboring tufted cells project to adjacent focal sites on the opposite side of the same bulb. Tufted cells comprising the IAS contain CCK and the terminals of these tufted cells synapse onto the dendrites of GABAergic granule cells (Liu and Shipley, 1994) The organization of IAS is reminiscent of the kind of point-to-point topography in other sensory pathways but the functional significance of this system is unclear. It might, however, further maintain the 'focal-diffuse' organization in the projection from the epithelium to the bulb (Adamek et al. 1986; Land, 1973; Land and Shepherd, 1974; Stewart, 1985). The focal organization of the IAS may be further preserved in at least one of the 510

The olfactory system

Ch. Ill

output pathways from the bulb originating in the AON, viz the pars externa system of the anterior olfactory nucleus (AONpE). The AONpE is a discrete architectonic subdivision of the AON. Mitral and tufted cells, including tufted cells giving rise to the IAS, project to AONpE (Schoenfeld and Macrides, 1984; Macrides et al. 1985). Axons of AONpE neurons project caudally, enter the anterior commissure, cross the midline and terminate in the contralateral olfactory bulb. The AONpE system terminates in the contralateral bulb in a modified point-to-point fashion. AONpE axons terminate with circumferential, but not longitudinal specificity, The AONpE projection terminates all along the longitudinal (rostrocaudal) axis of the bulb but is restricted to a limited band along the circumference of the bulb (Schoenfeld and Macrides, 1984). The topography in the combined intra- and interbulbar pathway is such that tufted cells associated with a focal part of the glomerular layer in one olfactory bulb project via the intrabulbar and interbulbar association pathways to a discrete region in the contralateral bulb (Macrides et al. 1985). However, the pE system terminates in a longitudinal (rostro-to-caudal) strip in the contralateral bulb. Taken together, these anatomical findings suggest that there may be epithelial to bulbar, intrabulbar and interbulbar pathways that are anatomically organized so as to preserve a plan of circumferential specificity in both the initial stages of the olfactory pathway and its connection to the contralateral bulb. The functional significance of this circumferential organization remains to be determined but it could be related to the recently discovered topographical segregation of different members of the olfactory receptor multi-gene family in the olfactory epithelium. These so-called 'expression zones' are organized as longitudinal strips in the olfactory epithelium. Such strips might project to longitudinal strips in the bulb. Connections of other subdivisions of A ON with MOB

In the rat approximately 60% of the non-pE AON neurons that project back to the ipsilateral bulb also project to the contralateral bulb (Alheid et al. 1984). These contralateral projections travel in the anterior commissure. The non-pE AON projections to the bulb do not appear to have any point-to-point or point/strip organization. They do, however, have some degree of radial organization. Axons of these AON neurons terminate in a laminar-like fashion at characteristic depths in the ipsi- and contralateral bulbs (Luskin and Price, 1983b). Most AON neurons project to the granule cell layer of the two bulbs but different AON divisions preferentially terminate in the superficial or deep part of the granule cell layer. This superficial-to-deep dimension represents the radial axis of the bulb and this axis is orthogonal to both the circumferential and longitudinal axis. Thus, the intrabulbar association pathway and the pE component of the interbulbar pathway have a degree of longitudinal and circumferential organization and the bulk of the AON interbulbar pathway and the centrifugal projections from AON back to the ipsilateral bulb have a degree of radial but, as yet, no suggestion of circumferential or longitudinal organization. The radial or laminar organization of the non-pE AON system may relate to a proposed plan of radial or laminar organization between mitral and tufted cells and granule cells. As reviewed earlier, the lateral dendrites of different mitral cells preferentially arborize at a particular depth of the EPL. The dendrites of some mitral cells arborize in the deeper parts of the EPL just above the mitral cell layer while the lateral dendrites of other mitral cells preferentially arborize more superficially in the EPL. The apical dendrites of granule cells located in the superficial part of the GCL arborize in 511

9

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TABLE 4.Neuropeptides in olfuctory cortical structures Neuropeptide

Location of cells

Cell SizeRype

Calbindin

AON-anterior, medial, lateral Taenia tecta- subpopulation of cells Piriform, entorhinal- mainly in layer 11, none in layer I

bi-multipolar

Cholecystokinin (CCK)

AON - anterior, dorsalis, lateralis, medialis Piriform cortex - mainly layer I1 Entorhinal cortex - layer I1 and 111

no information rare (21) rat rat ~yramidal/mult 12-17 rat no information cells/section no information

21 24 8

Corticotropin releasing factor (CRF)

AON-all subdivisions, Piriform-present in all layers

small

rat, monkey

3,18

Dynorphin B

Taenia tecta; AON-medial, ventral, posterior, dorsal; olfactory tubercle; piriform cortex, layer 11, layer 111, lat. entorhinal

non-pyramidal fusiform, scattered multipolar cells

rat

6

Enkephalin

ICC-AON, piriform-mainly layer I1 ISH-taenia tecta, AON (few), olf. tubercle, piriform (layer 11, some 111)

fusiform few pyramidatlmult varied

rat rat

16

LHRH

cells associated with medial olf. tract, olf. tubercle

1&15 p m

scattered

rat, others

2.25

GABNGAD

all olfactory regions

-small

rat

14

Piriform - layer la, Ib, 11. 111

-1 1-1 9 p m

-varying density -varying

Neuro tensin

piriform, entorhinal

no information

decrease w/age

rat

10

NPY

AON-all divisions, densest in medial, ventral and ventral-posterior subnuclei. - layers 11, 111, peduncular white matter Olf. tubercle - throughout Islands of Calleja - cells border the islands - within islands

-medium (1 5-25 pm) medium mediumlarge (> 25 pm)

- 2-5lsection - lO/section - IOlsection - llsection

rat

5

rat rat rat cat

5 5 5 19

human, rat

4,15

rat

4

Parvalbumin

AON-complementary to calbindin-IR neurons in AON: mainly in rostra1 part of ventral, lat. and dorsal subdivision, numerous in taenia tecta Piriform - mainly in layer 111

Cell number

Species

rat, human 4 8 % of cells

- multipolar

- dense

7,15 4

rat

varies with layer

Refs.

4

11

opossum

9

5

h

0

.=i TABLE 4. (coniinued)

~

_

_

_

22

~

SP

AON - ISH mRNA present in all subdivisions Olfactory tubercle Olfactory cortex - no ICC labelled cells present

small and medium

numerous sparse

VIP

Piriformientorhinal - mostly layer 11, some layer 111, feu in mol. layer

bipolar, 12 p n

scattered

-

~-

rat

23

rat, cat

12,17,22

rat, mouse, cat

1,13,17,20

-

References 1. Abrams, G.M., Nilaver, G. and Zimmerman, E.A. (1985); 2. Barry, J., Hoffman, G.E. and Wray, S. (1985); 3. Bassett, J.L., Shipley, M.T. and Foote, S.L. (1992); 4. Ceho, M.R. (1990); 5. De Quidt, M.E. and Emson. P.C. (1986); 6. Fallon, J.H. and Leslie, F.M. (1986); 7. Garcia-Segura, L.M., Baetens, D., Roth, J., Norman, A.W. and Orci, L. (1984); 8. Greenwood, R.S., Godar, S.E.. Reaves, Jr., T.A. and Hayward, J.N. (1981); 9. Haberly, L.B., Hansen, D.J., Feig, S.L. and Presto, S. (1987); 10. Hara, Y., Shiosaka, S., Senba, E., Sakanaka, M., Inagaki, S., Takagi, H., Kawai, Y., Takatsuki, K., Matsuzaki, T. and Tohyama, M. (1982); 11. Harlan, R.E., Shivers, B.D., Romano, G.J., Howells, R.D. and Pfaff, D.W. (1987); 12. Ljungdahl, A,, Hokfelt, T. and Nilsson, G. (1978); 13. Loren, I., Emson, P.C., Fahrenkrug, J., Bjorklund, A., Alumets, J., IIakanson, R. and Sundler, F. (1979); 14. Mugnaini, E. and Oertel, W.H. (1985); 15. Ohm, T.G., Muller, H. and Braak, E. (1991); 16. Petrusz, P., Merchenthaler, I. and Maderdrut, J.L. (1985); 17. Roberts, G.W., Woodhams, P.L., Polak, J.M. and Crow, T.J. (1982); 18. Sakanaka, M., Shibasaki, T. and Lederis, K. (1987); 19. Sanides Kohlrausch, C. and Wahle, P. (1990b); 20. Sanidcs Kohlrausch, C. and Wahle, P. (1990a); 21. Vanderhaeghen, J.J. (1985): 22. Wahle, P. and Sanides Kohlrausch, C. (1990); 23. Warden, M.K. and Young, W.S. (1988); 24. Westenbroek, R.E., Westrum, L.E., Hendrickson, A.E. amd Wu, J.-Y. (1987); 25. Zheng, L.M., Caldani, M. and Jourdan, F. (1988).

3

1

Ch. III

M.T. Shipley, J.H. McLean, L.A. Zimmer and M. Ennis

the superficial part of the EPL while the apical dendrites of granule cells located deeper in GCL preferentially arborize in the deeper parts of the EPL. Since the apical dendrites of granule cells have significant synaptic interactions with the lateral dendrites of mitral/ tufted cells, it is possible that superficial granule cells preferentially synapse with mitral cells whose lateral dendrites arborize in superficial EPL while deeper lying granule cells preferentially interact with mitral cells whose apical dendrites arborize in the deeper parts of the EPL. Mitral and tufted granule cell interactions therefore, may have a significant degree of radial organization. Since different divisions of the non-pE AON system terminate at different depths of the granule cell layer, there is a potential anatomical substrate to link, and possibly coordinate, mitral and granule cells with similar radial organization between the two olfactory bulbs. This radial organization may also differentially influence the outputs of the bulb as Scott and co-workers have shown that superficially located tufted cells tend to project preferentially to rostral parts of the olfactory cortex while deeper striatal, tufted and most mitral cells project preferentially to more caudal parts of the olfactory cortex (Scott et al. 1980; Scott, 1981; Schneider and Scott, 1983; Scott and Harrison, 1987). Much remains to be learned about the anatomy and the functional organization of MOB and AON circuitry. However, not withstanding the frequent statement that the olfactory system lacks topographical organization, it should not be inferred that this system is devoid of anatomical organization. The connections of different populations of MOB and AON neurons do express different degrees of circumferential, longitudinal and radial organization. An important problem for future research in olfactory neuroanatomy, therefore, is to learn if there are additional dimensions of organization and to determine how subpopulations of relatively odorant-specific ORNs articulate with the anatomical organization of the MOB, AON and olfactory cortex. 3.1.5. Transmitters of AON Candidate transmitters in AON and other olfactory cortical areas are summarized in Table 4. Aspartate has been proposed as a transmitter of AON neurons (mainly the dorsal and external divisions of AON) based on selective retrograde transport of 3H aspartate (Watanabe and Kawana, 1984; Fuller and Price, 1988). Fewer neurons in other subdivisions of AON contain aspartate, and no neurons in other known afferents to the bulb contain aspartate. There are a few met-enkephalin and somatostatinergic neurons in AON and some of these appear to project to the olfactory bulb (Davis et al. 1982). It appears that all neurons in AONpE contain the neuropeptide CRF (Shipley, in preparation). Besides the neurotransmitters/peptides discussed above, immunoreactive cells and fibers to calcium binding proteins have been described in AON. In this respect, calretinin immunoreactive fibers are found in the molecular layer o f A O N (Jacobowitz and Winsky, 1991), calbindin positive cells are found in all subdivisions of AON (Garcia-Ojeda et al. 1992; Celio, 1990) while parvalbumin positive cells are found in all subdivisions except for the medial one where the cells are quite sparse (GarciaOjeda et al. 1992; Celio, 1990). 3.1.6. Transmitter receptors in AON (Table 5) Like the MOB, the AON contains all mAChR's (ml-m4) and nAChR's (Hunt and Schmidt, 1978; Spencer et al. 1986; Fonseca et al. 1991; Rotter et al. 1979). Only the adrenergic beta l receptor shows intense staining in AON, while signals for beta2 and 514

The olfactory system

Ch. III

alpha2 receptors are similar to background levels (Wanaka et al. 1989; Palacios and Kuhar, 1982; Nicholas et al. 1993). Each glutamate receptor subtype listed is present in AON (Petralia et al. 1994; Petralia and Wenthold, 1992; Ohishi et al. 1993; Monaghan et al. 1985; Shigemoto et al. 1993; Martin et al. 1993; Gall et al. 1990; Molnar et al. 1993). While laminar organization is not noted, immunocytochemical evidence suggests that AMPA receptors are located to fusiform and large pyramidal cells in AON (Martin et al. 1993). In situ and autoradiographic studies suggest that GABA A receptors are present in higher amounts than GABAB receptors in AON (Zhang et al. 1991; Bowery et al. 1987; Richards et al. 1987; Palacios et al. 1981). Further study is needed to identify the laminar organization of these receptors. 3.1.7. Functions of AON Our understanding of the functional significance of AON is still rudimentary. Clearly, the major interbulbar connections of the AON implicate this structure in the interhemispheric processing of olfactory information. There is evidence that binasal mechanisms may function in spatial localization of odors (Bennett, 1968) and the AON system would be suspected to play a significant role in such mechanisms. Kucharski and Hall (1988)

T A B L E 5. Receptor subtypes of the A ON

Receptor

AON

References

mAChR1 mAChR2 mAChR3 mAChR4 nAChR

+ + + + +

2,4,15,17

+ + + + +

alphal

8,11,18

alpha2 betal beta2

+ + + + + +

Kainate

+ +

NMDA AMPA

Metabotropic

+ + + + + +

GABA A GABAB

+ + + +

3,5,7,9,12,13,16

1,10,14,19

Key: + +, receptors; - - , receptors absent; NR, not reported. Abbreviations: AON, anterior olfactory nucleus; mAChR, muscarinic cholinergic receptor; nAChR, nicotinic cholinergic receptor; alpha and beta adrenergic receptors; NMDA, N-methyl-D-Aspartate receptor; A M P A , ~-amino-hydroxy-5-methyl-4-isoxazolepropionic acid receptor; GABAAm, gamma aminobutyric acidreceptor. References 1. Bowery et al., 1987; 2. Fonseca et al., 1981; 3. Gall et al., 1990; 4. Hunt and Schmidt, 1978; 5. Martin et al., 1993; 6. Molnar et al., 1993; 7. Monaghan et al., 1985; 8. Nicholas et al., 1993; 9. Ohishi et al., 1993; 10. Palacios et al., 1981; 11. Palacios and Kuhar, 1982; 12. Petralia and Wenthold, 1992; 13. Petralia et al., 1994; 14. Richards et al., 1987; 15. Rotter et al., 1979; 16. Shigemoto et al., 1993; 17. Spencer et al., 1986; 18. W a n a k a et al., 1989; 19. Zhang et al., 1991.

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have shown that the AON is required to access and recall existing olfactory memories stored in the contralateral AON or olfactory bulb. 3.2. ROSTRAL OLFACTORY CORTEX Several small cortical areas located on the medial wall of the rostral hemisphere comprise part of the olfactory cortex that are often ignored by olfactory researchers, probably because of confusion about the cytoarchitecture and connections of these regions. These areas include the indusium griseum (or dorsal hippocampal continuation), the anterior hippocampal continuation (tt3) and taenia tecta (tt2). Recently, anatomical studies have begun to unravel the cytoarchitecture and connections of these regions and of the infralimbic cortex which may have interesting olfactory and visceral integration functions. 3.2.1. Indusium griseum

The indusium griseum (IG) or dorsal hippocampal continuation receives input from but does not project to the olfactory bulb. It is a thin layer of cortex which runs parasagittally just dorsal to the corpus callosum. IG has been the subject of debate as to whether it is more related to the hippocampus or olfactory bulb (cf. Wyss and Sripanidkulchai, 1983; Adamek et al. 1984 for further discussion). It now seems clear that IG receives direct inputs to its tiny molecular layer from the olfactory bulb (Wyss and Sripanidkulchai, 1983; Adamek et al. 1984; De Olmos et al. 1978). This input is mainly to the rostral IG with fewer fibers running more caudally. The molecular layer of IG also receives input from the lateral and medial entorhinal cortex (Luskin and Price, 1983b). Since the entorhinal area receives direct olfactory bulb inputs and, in turn, projects to the dentate gyrus of the hippocampus it has been suggested that IG is a phylogenetically old part of the hippocampus that receives direct olfactory information as opposed to most of the hippocampus that receives only indirect olfactory input via the entorhinal area (Adamek et al. 1984). 3.2.2. Anterior hippocampal continuation

The anterior hippocampal continuation (AHC) has been described in detail elsewhere (Wyss and Sripanidkulchai, 1983; Adamek et al. 1984). It is located just ventral to the rostrum of the corpus callosum and dorsal to the taenia tecta. The AHC has also been called tt3 (Switzer et al. 1985) and has been described in detail in the rat (Wyss and Sripanidkulchai, 1983). The inputs to AHC are similar to IG as are its efferent connections with the outstanding difference that IG does not project to the olfactory bulb while there is a modest projection from AHC to the olfactory bulb (Wyss and Sripanidkulchai, 1983; Scheibel and Scheibel, 1978; De Olmos et al. 1978). Other major efferent projections of the IG and AHC are to the mammillary bodies and anterior thalamic nuclei. 3.2.3. Taenia tecta

The taenia tecta proper (tt2) projects strongly to the olfactory bulb (De Olmos et al. 1978; Shipley and Adamek, 1984). Haberly and Price (1978b) divided the taenia tecta into dorsal and ventral subdivisions. The ventral subdivision (Fig. 16) has reciprocal connections with the MOB and with parts of the olfactory cortex. The neurons of this cortical 516

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Fig. 16. Cytoarchitecture of medial prefrontal cortex. A. Photomicrograph (darkfield illumination) of a coronal section through prefrontal cortex processed for acetylcholinesterase (ACHE) histochemistry. B. Higher power micrograph (brightfield illumination) of a section adjacent to that shown in (A). Note dense reaction product in discrete subfields of medial prefrontal cortex. C and D. High power darkfield (C) and lightfield (D) micrographs of the same section shown in (A) reveal the pattern of AChE and Nissl staining, respectively. Arrowheads indicate boundaries of cortical regions delineated by AChE and Nissl staining. Structures between the arrowheads from dorsal to ventral are: prelimbic cortex, infralimbic cortex, dorsal peduncular cortex and tenia tecta.

s t r u c t u r e are relatively t i g h t l y p a c k e d a n d are l o c a t e d d o r s a l to the a n t e r i o r o l f a c t o r y nuclei o n the m e d i a l wall o f the h e m i s p h e r e a n t e r i o r to the r o s t r u m o f the c o r p u s callosurn.

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3.2.4. Infralimbic cortex

The infralimbic cortex is mentioned here because of its secondary connections from olfactory inputs and because of its extensive connections with autonomic brain centers. This cortical region is found slightly rostral to but in the same general region as the AHC (Figs. 8-10 and 16 (Paxinos and Watson, 1986)). Although most studies indicated a lack of direct connections between infralimbic cortex and the olfactory bulb (Shipley and Adamek, 1984; De Olmos et al. 1978; De Carlos et al. 1989), at least one study has reported a weak projection (Neafsey et al. 1986). The cells in the infralimbic cortex that are said to project to the bulb appear to be in the same region that projects to the visceral centers of the brain (Neafsey et al. 1986). The infralimbic cortex also has direct dense projections to the molecular and polymorph layers of rostral piriform cortex and, possibly, endopiriform cortex (Hurley et al. 1991). Thus, infralimbic cortex may be a linkage between olfaction and autonomic function. 3.2.5. Olfactory tubercle

The olfactory tubercle in rodents, rabbits and other macrosomatic mammals is a prominent bulge on the base of the hemisphere just caudal to the olfactory peduncle. In such species, axons ofmitral and tufted cells (Heimer, 1968; Price, 1973; De Olmos et al. 1978) terminate in the superficial layer of the tubercle as in AON and primary olfactory cortex. The tubercle has a superficial plexiform layer like AON and primary olfactory cortex but the cellular architecture of the tubercle is intermediate between a cortical and a striatal structure. Immediately deep to the plexiform layer is a layer of neurons with apical dendrites that extend into the plexiform layer. Neurons deep to this so-called cortical layer, however, are not like layer III pyramids of primary olfactory cortex but rather are polymorphic, and their dendrites do not appear to preferentially extend into the plexiform layer as those of the pyramidal cells of layer III in olfactory cortex. These polymorphic neurons appear to be more akin to neurons of the striatum and indeed, Heimer's extensive neuroanatomical analysis of the tubercle and adjacent basal telencephalic gray matter has led to the concept of the 'ventral striatum' (Heimer and Wilson, 1975). This concept is beyond the scope of this chapter but its definition is based on parallel patterns of cytoarchitecture, transmitters and connectivity of ventral parts of the striatum and the tubercle with the dorsal neostriatum. Heimer and co-workers conclude that the ventral striatum is anatomically similar to the more familiar dorsal striatum (including the caudo-putamen), except that whereas the extrinsic and subcortical affiliates of the dorsal striatum are connected with neocortex and associated parts of the intralaminar/thalamic nuclei those of the ventral striatum are connected with cortical regions of the limbic system and the mediodorsal thalamic nucleus. Consistent with the striatal motif of the tubercle, the part of this structure that receives olfactory bulb afferents is much reduced in humans and other microsmatic primates despite the continued absolute expansion of the tubercle. Viewed in this context, the prominent olfactory input to the tubercle of macrosmatic species may reflect an expanded role for olfactory influence in such species on ventral striatal functions which are thought to be more related to the emotional sphere than those of the dorsal striatum. The tubercle also differs from the POC in that it does not send a reciprocal projection back to the bulb. This generalization needs to be somewhat qualified because the projection from the magnocellular basal forebrain, the nucleus of the diagonal band (DB) which is the sole source of cholinergic innervation of the bulb (Macrides et al. 1981; Shipley 518

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et al. 1986), may be considered a kind of centrifugal return from the ventral striatum, but at present there is no evidence that the olfactory bulb projection to the plexiform layer of the tubercle has direct or indirect anatomical linkages to influence the NDB neurons. Thus, the tubercle differs from the rest of POC in that it does not contain a population of cortical neurons that reciprocate the projection from the bulb. 3.2.6. Nucleus of the lateral olfactory tract (NLOT)

The cytoarchitecture of NLOT (Fig. 17C) has been studied extensively by McDonald (1983). It is considered an anterior part of the amygdala. NLOT can be subdivided into 3 layers on the basis of Nissl preparations; a superficial plexiform layer I which contains a few small and medium-sized cells, a layer II which contains many tightly packed cells, and layer III located dorsal to layer II and containing fairly large, loosely packed cells. Most cells of NLOT are medium-sized pyramidal shaped with extensive spines on secondary and distal dendrites. According to McDonald (1983), layers I and II appear similar in connections to the piriform cortex while layer III seems to be a closely related subcortical area. Many neurons of layers II and fewer neurons of layer III project to the olfactory bulb (de Olmos et al. 1978; Shipley and Adamek, 1984). In addition to olfactory bulb projections, many axons of NLOT neurons make up the stria terminalis and cross to the contralateral piriform cortex, olfactory tubercle, lateral nucleus of the amygdala, and bed nucleus of the stria terminalis (de Olmos, 1972). Afferent connections to NLOT arise mainly from olfactory related areas and the basolateral nucleus of the amygdala. 3.3. LATERAL OLFACTORY CORTEX 3.3.1. Architecture of the lateral olfactory cortex

The caudolateral part of AON is continuous through transitional zones with the piriform cortex, which in turn gives way caudally to periamygdaloid and transition cortices and then the lateral entorhinal cortex. Collectively, these cortical structures comprise the entire temporal cortical mantle ventral to the rhinal sulcus. Piriform cortex (Fig. 15 C-D; 17 A-F), also referred to as pyriform or prepyriform cortex, is a phylogenetically old, paleocortical structure. PC is located along the entire length of the lateral olfactory tract at the ventrolateral convexity of the base of the cortex. PC is thicker and more elaborate caudally than it is rostrally. PC is allocortical, having only two-three cellular layers and is thinner and less complex than the neocortex which has six layers. Haberly and Price (1978b) divided the piriform cortex into 3 layers that are further subdivided on the basis of cytoarchitecture and afferent connections. Layer I, the molecular layer, is the most superficial layer of PC. This layer is densely synaptic. The superficial half of this layer (Ia) contains the axons and synaptic terminals of mitral/tufted cells. The deep half (Ib) contains axons and terminals from ipsi- and contralateral olfactory cortical association inputs. Layer II is a distinct, narrow, tightly packed cell layer containing pyramidal neurons whose apical dendrites extend into layer I. Some smaller cells in layer IIa lack basal dendrites (Haberly and Price, 1978b) and are reminiscent of dentate granule cells in the hippocampus. Layer III is a thicker but less densely packed cellular layer containing larger pyramidal cells whose dendrites also extend throughout layer I. Layer III also contains large multipolar cells whose dendrites do not enter layer I and other intrinsic neuronal types. Layer III exhibits a superficial 519

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9 Fig. 17. Cytoarchitecture of olfactory cortex. A-H. Nissl stained coronal sections through olfactory related structures, principally showing the piriform and entorhinal cortices and amygdala nuclei. Bar in H, 1 mm.

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to deep pyramidal cell gradient with a higher density of pyramidal cells superficially. The deepest part of PC has been referred to by some as layer IV and by others as the endopiriform nucleus; this structure was recognized by Loo (1931) in the opossum and was further described by Haberly and Price (1978b). The cells of the endopiriform nucleus are mainly multipolar and are morphologically similar, but more densely packed than multipolar cells in layer III of PC. Multipolar cells and the deep layer III pyramids give rise to the subcortical outputs of PC. Layer II and superficial layer III pyramids project to the bulb (centrifugal fibers) and to other rostral and caudal levels of PC (association fibers). PC also contains numerous interneurons. The distribution of GABAergic interneurons has been described for the opossum and appears to be quite similar in the rat (Haberly, personal communication). These cells are found in all layers of piriform cortex (Fig. 25), including layer I, where they may function as a feedforward inhibitory system. There have been numerous reports of neuropeptide containing neurons in olfactory cortex; many of these have a morphology consistent with an interneuron, but little is known about their connections or functions. As noted by de Olmos et al. (1978), there are several transitional regions (including periamygdaloid cortex) between the piriform cortex and the entorhinal cortex and between olfactory cortices and neocortex. These transitional regions have been described in detail previously (De Olmos et al. 1978) and the reader is referred to that paper for additional information. Caudal to the piriform and periamygdaloid cortices is the entorhinal cortex. This cortex is divided into medial, lateral and intermediate divisions and has six layers. 3.3.2. Neuron types in the piriform cortex

There have been surprisingly few detailed studies of the morphology and distribution of PC neuronal types. The primary study cited in this regard was conducted by Haberly (Haberly, 1983) in the opossum. However, neuronal types and interconnections in the opossum are similar to those in rat (Haberly, 1983; Haberly and Behan, 1983; Haberly et al. 1987; Haberly, personal communication). Pyramidal cells

PC has two principal layers of pyramidal neurons, layers II and III, corresponding to superficial and deep pyramidal cells (Fig. 26). A third morphologically distinct subtype, the middle pyramidal cell has also been suggested, but is supported by only limited evidence (Martinez et al. 1987). Pyramidal neurons have several characteristic features and are similar to those in other cortical regions and the hippocampus: (1) A primary apical dendritic trunk that extends radially towards the pial surface and arborizes into numerous smaller branches that ramify in layer Ia and Ib (Haberly, 1983; Martinez et al. 1987). Some of these branches turn and run parallel to the pial surface for short distances. (2) A large number of relatively thin secondary or basal dendrites that emerge from the soma and extend several hundred microns into deeper parts of PC. Both the apical and basal dendritic tree are heavily invested with spines and varicosities. (3) A myelinated axon that typically extends deep to the soma terminates on other local pyramidal cells and interneurons (see below). Superficial pyramidal cells located in the external part of layer II, have somata of approximately 17 r in diameter. The apical dendrite of these neurons is relatively short 522

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and thick. Deep pyramidal cells are slightly larger than superficial pyramidal cell (mean diameter = 20 r and their apical dendrites are longer, typically extending unbranched from layer III to layer I. The collaterals of deep pyramidal axons travel into the endopiriform nucleus (Tseng and Haberly, 1989) whereas those of superficial pyramidal neurons do not appear to enter this structure. The apical dendritic tree of the superficial pyramidal generally branches more extensively and is more densely invested with spines and varicosities of deep pyramidal neurons. Although the morphology of superficial and deep pyramidal cells is similar, as discussed below, the neurophysiological properties of these two neuronal classes are somewhat different.

Intrinsic neurons There are a number of distinct intrinsic or interneuronal subtypes that differ in size and shape of the soma and dendritic organization.

Layer I." Horizontal, spiny, smooth and neurogliaform cells Horizontal cells are distributed almost exclusively in the superficial part of layer I (Haberly, 1983), although a cell type with a similar morphology has been described in layer III (Martinez et al. 1987). This cell type is characterized by a large fusiform soma of approximately 25 r in diameter that is oriented horizontally to the pial surface. The soma has a distinctive spiny appearance due to a large number of protuberances or knobs. These specialized appendages often extend from the soma on slender stalks. The dendrites of horizontal cells are oriented parallel to the pial surface and ramify within layer IA, and to a lesser extent, in layer Ib. A population of layer Ia horizontal cell with similar morphological characteristics stain for GAD, suggesting that they are GABAergic (Haberly et al. 1987). The spiny cell is a medium sized neuron (15-25/.tm in diameter) that is located throughout layer I. These neurons are characterized by small spines on the distal and proximal dendritic segments. Unlike the horizontal cell, spiny cells lack knobs or similar appendages on the soma. Spiny cell dendrites extend into all parts of layer I and less frequently into deeper layers. The spiny neurons gives rise to an unmyelinated axon that is horizontally oriented. Smooth cells present in layer I exhibit a variety of cell shapes and sizes and are distinguished by a lack of spines on the cell body and dendrites and the presence of beaded varicosities on the distal dendrite. Smaller sized smooth cells have a distinctive appearance that resembles neurogliaform cells present in some subcortical areas. The neurogliaform cell body is spherical and gives rise to thick dendritic trunks that branch, often at right angles, into a number of thin beaded fibers; this dendritic tree is usually restricted to a single sublamina in layer I. This neurogliaform cell type is located throughout layer I and is the most common cell type in this layer. The axons of neurogliaform cells are difficult to stain, but are unmyelinated and branch extensively. Layer II." Semilunar and neurogliaform cells The semilunar neuron is located in layer IIa and lacks a basal dendritic tree. This cell type has several apical dendrites that emerge at an oblique angle and arborize in layer I, and an axon that extends deep to the soma. The dendritic spines on this cell type are comparatively larger and less numerous than those on the pyramidal cell, and are covered with relatively large spines. Neurogliaform cells are similar to those described above. 523

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Layer IlL" Smooth and neurogliaform, multipolar complex appendage and giant cells The most numerous non-pyramidal cell in layer III are neurons that have relatively few dendritic spines or knobs. These cells are distributed in the middle and deep parts of layer III. The somata and dendrites of this cell type vary considerably. The dendrites emerge from numerous sites on the soma and usually branch once or twice near the soma and then extend radially in all directions with few subsequent bifurcations. The dendritic tree typically respects the boundaries of layer III. The axons of layer III smooth cells are difficult to stain, in contrast to layer I smooth cells, possibly because of myelination. The multipolar cell is located uniformly throughout layer III and the endopiriform nucleus (Tseng and Haberly, 1989). Multipolar cells give rise to a large number of branched, spiny dendrites that emerge from many sites on the cell body radiate in all directions, but are confined to layer III. Axons originate from the soma or proximal dendritic trunks and collateralize extensively within layer III, forming boutons en passant and boutons terminaux. Although axons of multipolar neurons have not been traced to layer I-II, these cells can occasionally be antidromically driven by layer I stimulation. Immunocytochemical studies indicate that many multipolar neurons are GABAergic, suggesting that they are inhibitory interneurons (Haberly et al. 1987). Haberly (1983) described a distinctive spiny neuron in layer III in the opossum. The cell body is typically spherical or fusiform and can exhibit spines. The distal dendrites of this cell type exhibit large, complex swellings that are connected to the dendritic trunk by small to large sized stalks. The appendages contain 10 or more individual swellings or knobs. The giant cell is a second category of spiny neuron present in layer III. The cell body is usually multipolar and as the name suggests, the size of the soma (mean diameter = 35/Ira) is the largest in PC. The giant cell is also typified by very large dendritic trunks that radiate in all directions, branch extensively, often at right angles, and then rapidly taper into slender distal fibers. While other neuronal types in PC are distributed throughout a particular layer, giant neurons are concentrated in the ventral parts of layer III. 3.3.3. Connections of the lateral olfactory cortex

Inputs from the olfactory bulb The main olfactory bulb sends a projection to the entire extent of piriform, periamygdaloid and lateral entorhinal cortex (see above, Outputs of MOB). This projection terminates in the superficial half of layer I, layer Ia. Both mitral and tufted cells project to the rostral parts of AON and piriform cortex while the projection to more caudal parts of olfactory cortex becomes progressively dominated by mitral cells (Schoenfeld and Macrides, 1984).

Feedback to the olfactory bulb Piriform cortex, lateral entorhinal cortex and the transitional cortical areas project heavily back to the olfactory bulb (Figs. 13, 14, 18, 19). The projections are heavier from the rostral than the caudal parts of primary olfactory cortex in rat and mouse (Shipley and Adamek, 1984). A few cells in the posterolateral and medial cortical amygdaloid areas may project to the MOB (Shipley and Adamek, 1984). These feedback projections to the olfactory bulb arise mainly from pyramidal neurons in layers II and III in primary olfactory cortex. 524

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Fig. 18. Connections of the MOB. A-E. A rostral to caudal series of coronal sections showing the patterns of anterograde and retrograde labeling produced by an injection of WGA-HRP in the main olfactory bulb.

The transmitter(s) of these olfactory cortical projections to the bulb are not known although glutamate is suspected because that excitatory amino acid is found in many cells in layers II and Ill of piriform and entorhinal cortex (Kaneko and Mizuno, 1988) (Table 4). These feedback projections from olfactory cortex to the olfactory bulb are believed to primarily excite the GABAergic granule cells in MOB which in turn inhibit firing of mitral cells (Nicoll, 1971) via dendrodendritic synapses between granule and mitral cell dendrites (Halasz and Shepherd, 1983). 525

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In the hamster, neurons in the rostral to caudal levels of piriform cortex terminate from superficial to deep in the GCL of MOB, respectively. However, this gradual shift in termination is not as apparent in the rat. The periamygdaloid cortex and NLOT terminate in the deep GCL (Luskin and Price, 1983). In summary, it would appear that most of the afferent projections to MOB from the AON project most heavily to the ipsilateral GCL. Intrinsic and association connections

In addition to the feedback projections to the olfactory bulb, olfactory cortex has other extensive connections which can be discussed as four classes: intrinsic or l o c a l - short connections between neurons in different layers of POC; associative - connections with different parts of POC, extrinsic - connections with other structures, and modulatory inputs - afferents that terminate in POC as part of a broader innervation of other cortical and subcortical neural systems. Intrinsic or local connections

There are extensive translaminar or local connections among PC neurons. Layer II pyramidal neurons give rise to extensive axon collaterals that innervate deeper layer III pyramidal cells and local inhibitory interneurons in layers I and II. The primary axons of pyramidal cells are myelinated and branch into a large number of stereotypical radiating, fine caliper unmyelinated collaterals (Haberly and Presto, 1986). The vast majority of axon collaterals are confined to layer III, although axons can be tracer into layer Ib. EM studies demonstrate that these collaterals give rise to synapses that terminate on dendritic spines and shafts of adjacent pyramidal cells as well as on the dendrites of deeper non-pyramidal GABAergic interneurons. Pyramidal cell axons are contacted by olfactory bulb terminals and by local collaterals of pyramidal cells. Deeper pyramidal cells also give rise to extensive local collaterals that may synapse with local interneurons or with more superficial pyramidal cells. Thus, there are extensive translaminar connections both from superficial to deeper layers and vice versa. In addition there are several classes of GABAergic and neuropeptide-containing neurons in PC and, although the connections of these neurons are not known, most of them have the appearance of local interneurons. GABAergic neurons appear to play an extremely important role in regulating olfactory cortical functions. Recent studies by Haberly's group indicate that intrinsic GABAergic cells may regulate LTP and the expression of epilepsy (see below). GABAergic neurons are scattered throughout layers II and III of PC (Fig. 25) and appear to regulate the excitability of the pyramidal cells via feedback inhibition. In addition, there is a prominent population of GABAergic neurons in layer I of PC. These cells may be excited directly by olfactory bulb inputs and possibly by association projections (see below) and function to regulate olfactory cortical excitability by feedforward inhibition Associational and commissural circuits

Haberly, Price and others have systematically studied the organization of intra-PC association and contralateral PC commissural circuits. This associational circuitry is extensive and exhibits a considerable degree of laminar and topographic specificity. These connections arise from pyramidal neurons in layer II and III. Neurons in layer 526

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IIa, IIb and III each give rise to discrete projections. Layer IIa neurons project to both more rostral and caudal portions of PC. Projections from layer IIb neurons are primarily caudally directed while those from layer III cell are predominantly rostrally directed (Haberly and Price, 1978a,b). PC association projections terminate exclusively in layer Ib, and thus, distribute in a non-overlapping, but complimentary manner to MOB inputs to the more superficial layer Ia (Haberly and Price, 1978a,b; Haberly, 1983). Association projections also exhibit a moderate degree of rostrocaudal polarity; projections from caudal PC to more rostral regions are much heavier than corresponding projections from rostral PC to more caudal regions. Projections from rostral and caudal PC also terminate with intralaminar specificity; rostral PC projections terminate heavily in the superficial part of layer Ib while projections from caudal PC terminate uniformly throughout layer Ib (Luskin and Price, 1983a,b). On the basis of these patterns of associational projection terminations, as well as termination patterns of other olfactory cortical structures, Luskin and Price (1983a,b) have defined two general classes of cortico-cortico projections: those terminating in layer IIb (the 'layer IIb fiber system') and those terminating in layer II and the deep part of layer Ib (the 'layer II-deep Ib fiber system). Commissural projections to the contralateral PC originate nearly exclusively from layer II neurons and travel in the anterior commissure [AC]. These projections innervate more posterior parts of the contralateral PC as well as nearby olfactory cortical sites (periamygdaloid cortex, lateral entorhinal cortex, anterior cortical nucleus, nucleus of the lateral olfactory tract) (Haberly and Price, 1978a,b). The caudally-directed commissural projections arise almost entirely from rostral layer IIb neurons. However, there are shorter, less extensive commissural projections from caudal PC that target rostrally adjacent regions. This pathway arises mostly from deep layer III neurons although there is a modest contribution from layer II neurons.

Physiology of commissural and ipsilateral association systems High frequency stimulation of AC depresses odor-induced activity in the bulb; section of AC augments this activity (Kerr and Hagbarth, 1955; von Baumgarten et al. 1962); however, section of AC reduces olfactory acuity (Bennett, 1968). Application of an odor to one epithelium a few msec before application of the same odor to the contralateral epithelium is reported to suppress a response recorded across the glomerular layer of the contralateral bulb (Leveteau and MacLeod, 1969). Both AC and piriform cortex (PC) stimulation produce a negative field potential recorded in the GCL (Walsh, 1959; Nakashima et al. 1978), as expected if excitatory currents are flowing into granule cells in that layer. Intracellular recordings from mitral cells (Yamamoto et al. 1963; Mori and Takagi, 1978b) demonstrate that AC stimulation produces an IPSP in mitral cells which has an appreciably slower rise time and longer duration than the IPSP generated by antidromic stimulation of LOT. These results indicate that AC and PC stimulation causes excitation of granule cells which release GABA onto mitral cells, resulting in an IPSP in the mitral cells. The response to AC stimulation summates with repeated shocks: single shocks produced no visible IPSP, but three or more shocks at 100 Hz caused a significant hyperpolarization accompanied by block of antidromic invasion of the impaled cell. Preceding AC stimulation blocks the IPSP which normally follows antidromic stimulation of LOT (Yamamoto et al. 1962), suggesting that the commissural system is able to modulate the strength of lateral inhibition in the mitral/granule system. 527

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Extrinsic outputs of olfactory cortex Two classes of POC outputs were discussed above- the feedback projection back to the olfactory bulb and the association connections between rostral and caudal olfactory cortex. A third class of outputs is treated separately because it represents the projections of POC to brain regions not generally included in the olfactory system per se although their receipt of inputs from POC obviously implicates these POC targets in olfactory function. The extrinsic outputs of POC are both to cortical and subcortical structures (Fig. 19). Neocortical projections In the mouse and rat the projection of the olfactory bulb to POC extends dorsally beyond the cytoarchitectural limits of POC into the ventral parts of the granular insular and perirhinal cortices (Shipley and Geinesman, 1984). Insular cortex in rodents contains the cortical representation of ascending pathways arising in the nucleus of the solitary tract (NTS) in the medulla (Shipley, 1982). NTS is the initial subcortical relay for gustatory and visceral sensory input to the CNS. The representation of NTS in insular cortex appears to comprise both a primary sensory cortical map for gustation and visceral sensation and also, via descending corticofugal projections, a route whereby cortex can modify visceral-autonomic and possibly gustatory function (Shipley, 1982). Thus, the direct olfactory bulb projection into ventral insular cortex in rodents has been suggested as one relatively direct route for the integration of olfactory and gustatory information in the neural representation of flavor and the integration of olfactory and autonomic information (Shipley and Geinesman, 1984). The existence of direct projections from the olfactory bulb to homologous insular cortical areas in the primate have not been established with anatomical methods. In rodents there are also direct projections from POC to the dorsally adjacent insular cortical fields involved in gustation and visceral sensation (Price, 1985; Shipley, unpublished observations) and there are projections from mediodorsal thalamus to insular cortex and to medial cortical fields that also preferentially project to hypothalamic and brainstem regions involved in autonomic function (Price, 1985). Thus, there are direct POC projections to gustatory-autonomic cortical areas and, though less well characterized, potential circuitry from POC to mediodorsal-submedial thalamus to the lateral and medial neocortical areas involved in gustatory and autonomic function. The precise homologies among the lateral and medial cortical fields linking olfactory with gustatory-autonomic systems and the corresponding fields in the primate brain remain to be worked out. However, electrophysiological studies indicate that neurons in potentially homologous cortical areas in primates respond to odors with a higher degree of selectivity than neurons in either the olfactory bulb or POC (Tanaki et al. 1975; Takagi, 1986; Yarita et al. 1980). Thus, it may be that cortico-cortical and cortico-thalamo-cortical circuits from POC to neighboring cortical areas play a role in flavor perception and in linking olfactory stimuli to the hypothalamic-autonomic axis. Subcortical projections: hypothalamus and thalamus The heaviest and most direct projections to the hypothalamus derive from neurons in the deepest layers of piriform cortex and the anterior olfactory nucleus. These projections terminate most heavily in the lateral hypothalamic area (Price, 1985). Some poly528

The olfactory system

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morphic neurons of the olfactory tubercle and part of the AON (Price, 1985), also project to the hypothalamus. Olfactory-recipient parts of the cortical and medial amygdaloid nuclei also project to medial and anterior parts of the hypothalamus. In infraprimate species, anatomical experiments have demonstrated a strong projection from POC to the magnocellular, medial part of the mediodorsal thalamic nucleus and the submedial nucleus (nucleus gelatinosa) (Benjamin et al. 1982; Price and Slotnick, 1983). Retrograde tracing studies show that the olfactory cortico-thalamic projection arises from neurons in the deepest layer of piriform, periamygdala and entorhinal cortex and the polymorphic cell layer of the olfactory tubercle (Price and Slotnick, 1983). These projections have apparently not yet been established with anatomical methods in primates but neurophysiological studies in primates indicate that neurons in the magnocellular medial part of the mediodorsal and in the submedial thalamic nuclei are responsive to olfactory bulb stimulation and odors (Benjamin and Jackson, 1974; Russchen et al. 1987; Yarita et al. 1980). Thus a strong output from all parts of POC to the mediodorsal and submedial thalamic nuclei appears to be a fundamental feature of olfactory circuitry and represents a potentially important route for the dissemination of olfactory information to other cortical and subcortical areas. POC also receives extrinsic subcortical modulatory inputs; these are discussed in Section 6. 3.3.4. Transmitter receptors in the lateral olfactory cortex (Table 6)

It is believed that many of the neurons in PC receive EAA inputs either from the lateral olfactory tract and/or from cortico-cortico connections within PC. Furthermore, electrophysiological studies show a role for EAA receptors in PC. Recent autoradiographic, in situ, and immunocytochemical evidence suggests that layer II PC contains an extensive amount of AMPA and kainate receptor subtypes, while layers Ia and II stain for NMDA receptors (Petralia and Wenthold, 1994; Monaghan et al. 1985; Wisden and Seeburg, 1993; Gall et al. 1990; Petralia and Wenthold, 1992,; van den Pol et al. 1994; and Molnar et al. 1993). Further study of the cellular identification of these receptors within PC is necessary. PC also contains local GABAergic interneurons in layer Ia and deep to pyramidal cells of layer II. In situ and autoradiographic studies suggest that GABAA receptors are located in layers I and III of PC (Young and Kuhar, 1980b; Bowery et al. 1987; Palacios et al. 1981). Physiological studies suggest that GABA receptor activation may play a role in pyramidal cell activity in layer II. Further studies, though, are needed to determine the cell type and neuronal location of these receptors. While studies have shown intense staining for GABAA,there is only a very weak signal for the presence of GABAB receptors in PC (Bowery et al. 1987). 3.3.5. Piriform cortex is a seizurogenic focus

Considerable evidence, particularly in the last five years, has spotlighted PC as a key seizurogenic site in the cerebral cortex and dysfunction of this area may play an important role in temporal lobe epilepsy. Systemic injection of muscarinic receptor agonists or acetylcholinesterase inhibitors produces robust, sustained seizure activity in PC. Several recent studies have demonstrated that PC is the first cortical or subcortical forebrain structure to exhibit increased c-fos expression, a marker for elevated neuronal activity, after convulsive doses of pilocarpine or the irreversible acetylcholinesterase 529

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TABLE 6. Receptor subtypes of piriform cortex Receptor

PC

References

mAChR1 mAChR2 mAChR3 mAChR4 nAChR

+ + (I,II) ++ ++ + + (II) + + (II,III)

2,5,8,22,23,25,26,33

alphal alpha2

+ + (II)

15,17,24,27,29,31

betal beta2

+ + (II) + + (II)

D1 D2

+ + (II) + +

3,7,9

+ +

6,10,11,12,20,21

5-HTIA 5-HTE~aC Kainate NMDA AMPA Metabotropic GABAA GABAB

+ + (II)

+ +/+ + (II) + + (II)

4,14,15,18,19,30,35

+ + (Ia,II) + + (II) NR + + (I,III)

1,16,32

+ +

Key: + +, receptors present; - - , receptors absent; NR, not reported Abbreviations: PC, piriform cortex; mAChR, muscarinic cholinergic receptor; nAChR, nicotinic cholinergic receptor; alpha and beta adrenergic receptors; D, dopamine receptors; 5-HT1A and 5-HTzA/o serotonin receptors; NMDA, N-methyl-D-Aspartate receptor; AMPA, ~-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid receptor; GABAA/B, gamma aminobutyric acidreceptor; XX, receptor present, receptor not present; NR, receptor not present or not studied. References 1. Bowery et al., 1987; 2. Buckley et al., 1988; 3. Fremeau et al., 1991; 4. Gall et al., 1990; 5. Hill et al., 1993; 6. Hoffman and Mezey, 1989; 7. Huang et al., 1992; 8. Levey et al., 1991; 9. Mansour et al., 1990; 10. McLean et al., 1995; 11. Mengod et al., 1990a; 12. Mengod et al., 1990b; 13. Molnar et al., 1993; 14. Monaghan et al., 1985; 15. Nicholas et al., 1993; 16. Palacios et al., 1981; 17. Palacios and Kuhar, 1982; 18. Petralia and Wenthold, 1992; 19. Petralia et al., 1994; 20. Pompeiano et al., 1992; 21. Pompeiano et al., 1994; 22. Rotter et al., 1979; 23. Sahin et al., 1992; 24. Sargent-Jones et al., 1985; 25. Segulla et al., 1993; 26. Spencer et al., 1986; 27. Unnerstall et al., 1984; 28. van den Pol et al., 1994; 29. Wanaka et al., 1989; 30. Wisden and Seeburg, 1993; 31. Young and Kuhar, 1980a; 32. Young and Kuhar, 1980b; 33. Zilles et al., 1989.

i n h i b i t o r s o m a n . M i c r o i n j e c t i o n s o f a n u m b e r o f n e u r o a c t i v e c o m p o u n d s unilaterally into the d e e p p a r t s o f P C (layers II-III) p r o d u c e s seizures w h i c h s p r e a d to the rest o f the b r a i n w i t h i n several m i n u t e s ( P i r e d d a a n d Gale, 1985; P i r e d d a a n d Gale, 1986). T h e m o s t a n t e r i o r p a r t of PC, p r e p i r i f o r m cortex, recently d e s i g n a t e d a r e a t e m p e s t a s ( H a l o n e n et al. 1994), is especially seizurogenic. F o c a l m i c r o i n j e c t i o n s o f the bicuculline, e x c i t a t o r y a m i n o acids ( E A A s ) or c a r b a c h o l into this restricted p a r t o f rostral P C initiate seizures w h i c h p r o p a g a t e t h r o u g h o u t the cerebral c o r t e x a n d to s u b c o r t i c a l limbic structures. T h e doses o f agents r e q u i r e d to initiate seizures was 20 to 30 times lower t h a n t h o s e n e c e s s a r y to initiate generalized seizures in a d j a c e n t f o r e b r a i n s t r u c t u r e s including the a m y g d a l a a n d h i p p o c a m p u s . M i l l a n et al. (1988) h a v e s h o w n t h a t m i c r o i n j e c t i o n s 530

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of kainate or quisqualate into PC trigger seizures in animals given subconvulsive doses of pilocarpine. Gale and colleagues (Halonen et al. 1994) have recently identified key relay sites involved in the spread of seizures triggered in deep prepyriform cortex to other forebrain structures. These studies found that microinjections of muscimol or non-NMDA receptor antagonists into posterior PC or the dorsally adjacent perirhinal cortex prevented the propagation of seizures from rostral PC. The role of NMDA receptors in seizure generation in PC is less clear. Microinjection of an NMDA receptor antagonist into PC was reported to prevent seizures induced by systemic injection of pilocarpine (Millan et al. 1986). However, later studies by the same group (Millan et al. 1988) found that microinjection of NMDA into PC did not increase seizure susceptibility to subconvulsive doses of pilocarpine and, paradoxically, prevented seizures induced by a convulsive dose of pilocarpine. Interestingly, intact afferent input to PC from the olfactory bulb appears to be required for the initiation of seizures in PC. Olfactory bulb deafferentation (bulbectomy) protects against pilocarpine-induced seizures and increases the dose of KA infused into PC necessary to produce seizures (Millan et al. 1988). Additional evidence pinpointing PC as a key site of seizure initiation is provided by kindling studies. Repeated, intermittent stimulation of the olfactory bulb (Russel and Stripling, 1985; Haberly and Sutula, 1992) and other limbic structures (McIntyre and Wong, 1986) readily trigger epileptiform afterdischarges in PC. This same phenomenon has been demonstrated in the hippocampus, amygdala and other cortical regions following stimulation of a number of brain sites. However, epileptiform discharges occur in PC after fewer kindling trials than in the hippocampus and subcortical limbic structures (Kairiss et al. 1984; Racine et al. 1988). In addition, seizure activity occurs in PC slices taken from kindled animals when superfused in standard Krebs solution, whereas similar activity is only observed in hippocampal slices from kindled animals when the potassium concentration is elevated or GABAergic inhibition is blocked (Kairiss et al. 1984; King et al. 1985). Recent in vitro studies from Haberly's laboratory suggest that deep layer III pyramidal cells and neurons in the endopiriform nucleus, considered by some as layer IV PC, may generate the initial epileptiform events that trigger seizures in PC. Following bursting evoked by 2-20 rain of low intensity shocks to layers I-III, high amplitude, long latency depolarizing potentials characteristic of epileptiform discharges are observed in pyramidal cells (Hoffman and Haberly, 1991). While these discharges were observed in both superficial and deep pyramidal cells, discharges were of greater amplitude in deep vs. superficial cells and only depolarizing potentials in the deep pyramidal cells triggered action potentials. In agreement with the in vivo studies of Piredda and Gale (1985, 1986), microapplication of high potassium, glutamate or cobalt after association-induced bursting elicited epileptiform discharges only when applied to deep PC and the endopiriform nucleus. Using an elegant microslice surgical dissection technique, this same study demonstrated that epileptiform discharges could be evoked in deep pyramidal cells isolated from the superficial layers of PC. These and other findings suggest that epileptiform discharges initiate in deep layer III PC and the endopiriform nucleus. Taken together, these studies indicate that PC is an extremely seizure susceptible area. Elevated neuronal activity in the rostral part of PC initiates seizures which propagate throughout the brain. As reviewed above, the activity of PC output neurons, pyramidal cells, appears to be tightly regulated and finely tuned by opposing neural circuits: extrinsic and intrinsic EAA circuits and interneuronal inhibitory GABAergic circuits. 531

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Relatively small disruptions in the balance between these excitatory and inhibitory influences appears to be sufficient to cause a focus of epileptiform activity in PC that spreads throughout the brain. In addition, we and others have shown that some of the modulatory transmitter inputs (acetylcholine) increase PC neuronal excitability and initiate seizures. It is not known how foci of epileptiform activity in PC becomes sufficiently intense to produce prolonged, sustained seizures (i.e., status epilepticus) that spread to other cortical and subcortical structures. Based on our work with cholinolytic seizures and other recent findings on the cellular actions of neurotransmitters, we have suggested that cholinergic overstimulation of PC neurons, for example, causes a depression of cellular regulatory mechanisms that normally limit the responsiveness of PC neurons to EAAs (E1-Etri et al. 1992). There is a well-established cellular mechanism by which ACh increases the excitability of cortical neurons, including PC neurons, to EAAs. ACh, acting via muscarinic receptor-mediated second messenger systems, potently blocks a Ca+-modulated, K+-mediated voltage dependent afterhyperpolarization which normally functions to prevent cortical neurons from being overexcited by EAAs. As a result, cholinergic stimulation of PC neurons may dramatically increase the responsiveness of these neurons to EAAs that are being tonically released by the majority of synapses impinging on them. Because these cortical neurons are progressively excited by EAA inputs, they release more EAAs at their own synaptic terminals on other cortical neurons, thus further feeding an EAA 'chain reaction'. This excess stimulation may eventually lead to potentiation of N M D A receptors, which further strengthens EAA synaptic connections. Sparenborg et al. (1990) have shown that the N M D A receptor antagonist, MK801, can prevent cholinolytic seizures, thus a role for EAA overstimulation has already been suggested. With sufficient strengthening of these synapses, the mutual excitation between EAA synapses may become self-sustaining and progressively lead to a positive feedback and feedforward stimulation in which normal EAA synaptic pathways between cortical neurons are able to sustain seizures. It is important to note, however, that events similar to those described above could be triggered directly as a result of imbalances in EAA and/or GABAergic synaptic transmission in PC. 3.3.6. Modeling of olfactory network function PC's extensive network of associative connections, taken with the distributed nature of afferent inputs to PC from the bulb has been modeled recently by Haberly, Bower and others as a distributed association network that is hypothesized to function as a content addressable memory for spatially distributed odor patterns.

4. INTEGRATION OF THE MAIN OLFACTORY SYSTEM WITH OTHER FUNCTIONS 4.1. ODORS AND COGNITION There are several sites where olfactory discrimination and cognition could arise as virtually all of the primary olfactory regions are cortical structures including the olfactory bulb, anterior olfactory cortex, piriform cortex and entorhinal cortex. In addition, olfactory information is routed to the neocortex via the thalamus (Fig. 19). Physiological studies in monkeys suggest that some degree of odor discrimination may take place in 532

The olfactory system

Ch. III OLFACTORYCORTEX!

CIRCULATION

l

PARASYMPATHETIC

SYMPATHETIC

Fig. 19. Some of the higher order connections of the main olfactory system. Emphasis is on possible circuits that mediate o u t p u t responses such as a u t o n o m i c or h o r m o n a l changes. O u t p u t projections of the M O B are shown as thick lines; higher order connections are shown as thin lines. Cortical structures are depicted as boxes; subcortical structures as ellipses. Abbreviations: A C o = anterior cortical a m y g d a l o i d nucleus; A O B = accessory olfactory bulb; A O N - anterior olfactory nucleus (m - medial division); BST - bed nucleus of the stria terminalis; Ce = central nucleus of the amygdala; D H R - dorsal h i p p o c a m p a l rudiment; D P C - dorsal peduncular cortex; D R - dorsal raphe nucleus; Ent - entorhinal cortex; H C - hippocamus; IC - insular cortex; I M L - intermediolateral cell column of the thoracic level of the spinal cord; L C - locus coeruleus; L P O = lateral preoptic area; M e = medial a m y g d a l o i d nucleus; M D = medial dorsal nucleus; M O B - main olfactory bulb; M R - median raphe; N A O T = nucleus of the accessory olfactory tract; N L O T = nucleus of the lateral olfactory tract; DB - nucleus of the diagonal band; O F C - orbital frontal cortex; PAC = periamygdaloid cortex; P A G = midbrain periaquaductal gray; P C o = posterior cortical a m y g d a l o i d nucleus; Pit - pituitary; Pir - piriform cortex; p N A / N A = nucleus ambiguus and periambiguual area; R V L = rostrventrolateral medulla; SO = supraoptic nucleus; Tu = olfactory tubercle; T T - taenia tecta. 10 - dorsal m o t o r nucleus of the vagus.

the lateral and posterior orbitofrontal cortex (see Takagi, 1984, for review). This olfactory information is relayed either through the mediodorsal thalamus or through corticocortical routes (Takagi, 1984). There are several studies showing a potential olfactoneocortical circuit via the thalamus. For example, the olfactory tubercle, insular cortex and PC receive input from mitral cell axons (Broadwell, 1975b; De Olmos et al. 1978; Ojima et al. 1984; Shipley and Adamek, 1984). The olfactory tubercle and PC project to the dorsomedial thalamic nucleus (Benjamin et al. 1982; Powell et al. 1965; Price, 1985) and the submedial thalamic nucleus (Price and Slotnick, 1983) although the projection from PC (but not insular cortex) has recently been questioned (Motokizawa 533

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M.T. Shipley, J.H. McLean, L.A. Zimmer and M. Ennis

et al. 1988). The dorsomedial thalamus projects to the posterior orbitofrontal cortex so this pathway may mediate some aspects of olfactory discrimination. Physiological evidence, however, suggests that the dorsomedial thalamus projection is stronger to the centroposterior portion of orbitofrontal cortex which, according to Takagi (1984), is more involved in integrating odor sensations than discriminating odors because individual cells in that region respond to several odors; single cells in lateral posterior orbitofrontal cortex more commonly respond to a single odor. Thus, cortico-cortical pathways may be involved in higher-order olfactory functions. The transmitters involved in the pathways discussed above are not known although one might suspect that the excitatory amino acid glutamate is involved since it is found in all thalamic nuclei and projection cells of piriform cortex (Kaneko and Mizuno, 1988). 4.2. OLFACTION AND TASTE/VISCERAL INTEGRATION Olfactory stimuli can activate visceral response and autonomic adjustments, such as gastric secretions, salivation, and changes in heart rate. The circuits that mediate these functions are becoming known (Fig. 19). One possibility is MOB and PC connections with the insular cortex (Fig. 19) might be involved in these functions (Saper, 1982; Shipley and Sanders, 1982; Shipley and Geinisman, 1984; Shipley, 1982; Ruggiero et al. 1987). Studies in the mouse and rat show that a portion of the granular insular cortex is a site of significant overlap between olfactory and visceral information (Shipley and Geinisman, 1984, mouse; Krushel and Van Der Kooy, 1988, rat). In addition, the medial frontal cortex may be an area of motor control of visceral activity (Neafsey et al. 1986). There are direct projections from MOB to the ventral part of the medial frontal cortex and there are reciprocal connections between insular and the medial frontal cortex. Olfactory pathways may act through the insular cortex and medial frontal cortex to influence autonomic and visceral function via direct projections to cardiovascular regions of the ventral medulla and the solitary nucleus (Ruggiero et aI. 1987). Alternatively, the insular cortex may influence cardiovascular regions by less direct routes. For example, the central nucleus of the amygdala, which receives a dense projection from the insular cortex (Shipley and Sanders, 1982), projects to brainstem autonomic centers such as the periaqueductal gray and dorsal vagal complex (Hopkins and Holstege, 1978; Hopkins et al. 1981; Rizvi et al. 1991). Portions of the periaqueductal gray project to the ventral lateral medulla (Van Bockstaele et al. 1989) and may be involved in pressor and depressor responses of the cardiovascular system (Carrive et al. 1987). Another region that may be involved in the integration of various senses is the posterolateral orbitofrontal cortex. This cortex receives input from the insular cortex (Wiggins et al. 1987) and many neurons in the orbitofrontal cortex of the primate react to both taste and smell (Wiggins et al. 1988) and even visual inputs (Rolls, 1989). Thus, the orbitofrontal region may be an area where higher level integration of multiple sensory modes (taste, smell, vision) takes place. 4.3. OLFACTION AND MOTOR ACTIVITY As noted earlier, several pathways link olfactory related structures to what Heimer and Wilson (1975) have termed the ventral striatum. These connections are proposed to provide a means by which limbic (and possibly, olfactory) information are integrated with the motor control regions of the striatum so that visceral and somatic effectors may be controlled by these pathways (Newman and Winans, 1980a). Olfactory linkages to 534

The olfactory system

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the ventral striatum are mediated by parallel projections from MOB, AON and piriform cortex to the olfactory tubercle. Neurons in the olfactory tubercle and some in piriform cortex project to the nucleus accumbens (part of ventral striatum) which in turn projects to the ventral pallidum and substantia nigra, pars reticulata (Newman and Winans, 1980a). The neurochemistry of the accumbens/olfactory tubercle projections are only open to speculation at this time. It would be interesting if the neurochemical circuitry parallels the cortex (glutamate) ~ neostriatum (GABA, enkephalin, substance P) -~ palIidal (GABA) ~ thalamus (glutamate- ?) ~ cortex loop of the basal ganglia. The transmitters/peptides and precise circuitry of those pathways have recently been reviewed (Albin et al. 1989) but the equivalent relationships in the ventral striatal pathways remain to be determined.

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Fig. 20. Olfactory epithelium projections to the MOB. Photomicrographs of sagittal sections through the olfactory bulb in sections stained for Nissl (A) or with WGA HRP after injection of the tracer in the olfactory epithelium (B). Note that most of the olfactory bulb is comprised by the main olfactory system while a small portion of the dorsocaudal bulb is occupied by the accessory olfactory bulb in the rat. Note also in B that the WGA HRP did not transport to the glomeruli of AOB since the tracer did not gain access to the vomeronasal organ that is embedded in the nasal septum. Bar in B, 1 mm.

535

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M.T. Shipley, J.H. McLean, L.A. Zimmer and M. Ennis

Fig. 21. Architecture of the AOB. Photomicrographs through the accessory olfactory bulb in Nissl stained sections (A,B) and a section stained with cytochrome oxidase (CO) (C).

4.4. OLFACTION AND MEMORY The entorhinal cortex receives a substantial input from the MOB (Broadwell, 1975; De Olmos et al. 1978; Kosel et al. 1981; Shipley and Adamek, 1984). In turn, the medial and lateral entorhinal cortex projects to the dentate gyrus and CA fields of the hippocampus (Hjorth-Simonsen, 1972; Steward, 1976). Recent studies show that MOB projections to entorhinal cortex make direct contact with stellate cells located in layer II that in turn project via the perforant path to the hippocampus (Schwerdtfeger et al. 1990). In addition, piriform cortex has direct connections to the entorhinal cortex. Because the hippocampus is important in memory function, these olfactory-entorhinalhippocampal circuits may be important for establishment or recall olfactory memories formed or associated with other events.

5. THE ACCESSORY OLFACTORY SYSTEM 5.1. ACCESSORY OLFACTORY BULB The accessory olfactory bulb (AOB) has some similar cytoarchitectural features to the MOB, but is much smaller (Fig. 20, 21). The AOB is located at the caudal-dorsal end of MOB. The vomeronasal nerve transmits information from the vomeronasal organ (VNO) to the glomeruli of AOB. The AOB does not receive projections from the main 536

The olfactory system

Ch. III

olfactory epithelium (Fig. 20B) nor does the VNO project to the MOB. The glomerular layer in AOB (AGL) is less distinct than in the MOB because the AOB glomeruli are fewer and smaller. In addition, the periglomerular cells are far fewer than in the MOB with the result that the glomeruli are not so neatly delineated by a shell of cell bodies. However, glomerular structure can still be observed with stains which highlight axonal activity (Fig. 21C). The term 'periglomerular' is thus less appropriate in AOB than MOB because the few periglomerular cells tend to be located superficial or deep to the glomeruli rather than in the regions between the glomeruli. The external plexiform layer of AOB (AEPL) and mitral cell layer (AMCL) are also less distinct than the corresponding layers of MOB. The AOB internal plexiform layer (AIPL) is unremarkable and is situated between the mitral cell layer and lateral olfactory tract. The granule cell layer (AGCL) of AOB, located deep to the lateral olfactory tract, contains the same type of small cell as in MOB granule cell layer. Despite being called the mitral cell layer by many authors, the output cells in AOB are much more polymorphic than their counterparts in MOB (Takami and Graziadei, 1991). 5.1.1. Neurotransmitters in the AOB (Table 7) Based on retrograde transport of labeled amino acids, aspartate is suspected to be a transmitter of AOB output neurons. More AOB mitral cells appear to be aspartatergic than MOB mitral cells (Fuller and Price, 1988). Many mitral cells in AOB of the guinea pig contain neurotensin (Matsutani et al. 1989), while in rat, mitral cells transiently express substance P, but the expression in these output cells gradually diminishes after postnatal day 10. Interestingly, substance P-IR granule cells increase in number at the time when mitral cell expression is decreasing (Matsutani et al. 1988). The few 'periglomerular cells' in AOB are neurochemically different from those in MOB. The most obvious difference is the lack of dopaminergic periglomerular cells in AOB. Also lacking are the substance P containing external tufted cells that are abundant in MOB of some species (Baker, 1986). GABAergic periglomerular and granule cells are present in AOB

TABLE 7. Candidate transmitters & the A OB Transmitter/ peptide location

Cell type

Cell size

Cell number

Species

Refs

Aspartate DA GABA met-ENK Neurotensin Substance P

output juxtaglomerular cells PG and - granule cells GL output output output GOL - granule cells E P L - mitral or (Van Gehuchten)

medium small small small medium 10-15 r medium 9r 12 r

many rare many few many several many numerous few

rat many mouse rat guinea pig rat hamster rat, hamster cat

1 2,3 2 4,5 6 5 3 3,5 not 7 8

VIP

References 1. Fuller, T.A. and Price, J.L. (1988); 2. Baker, H., Towle, A.C. and Margolis, F.L. (I988); 3. Baker, H. (1986); 4. Gouda, M., Matsutani, S., Senba, E. and Tohyama, M. (1990); 5. Matsutani, S., Senba, E. and Tohyama, M. (1988); 6. Matsutani, S., Senba, E. and Tohyama, M. (1989); 7. Macrides, F. and Davis, B.J. (1983); 8. Sanides Kohlrausch, C. and Wahle, R (1990a).

537

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M.T. Shipley, J.H. McLean, L.A. Zimmer and M. Ennis

TABLE 8. Receptor subtypes of the A OB Receptor

AOB

mAChR nAChR

NR NR

alphal/2 betal/2

NR NR

D1 D2

NR NR

5-HT1A 5-HT2A/c

NR NR

Kainate NMDA AMPA Metabotropic

NR + + + + + +

2,4,5

GABAA GABAB

+ + NR

1,3,6,7,8

References

Key: + +, receptors present; - -, receptors absent; NR, not reported Abbreviations: AOB, accessory olfactory bulb; mAChR, muscarinic cholinergic receptor; nAChR, nicotinic cholinergic receptor; alpha and beta adrenergic receptors; D, dopamine receptors; 5-HT1A and 5-HTzA/c, serotonin receptors; NMDA, N-methyl-D-Aspartate receptor; AMPA, ~-amino-3-hydroxy-5-methyl-4isoxazolepropionic acid receptor; GABAA/B, gamma aminobutyric acid receptor; XX, receptor present in region. References 1. Laurie et al., 1992; 2. Ohishi et al., 1993; 3. Persohn et al., 1992; 4. Petralia and Wenthold, 1992; 5. Petralia et al., 1994; 6. Richards et al., 1987; 7. Young and Kuhar, 1980b; 8. Zhang et al., 1991.

(Baker et al. 1988). Substance P containing cells are most prominent in the AGCL of rats; in contrast, there are fewer substance P-IR cells in AGCL in rabbit, guinea pig, cat and hamster and in mice these cells appear to be absent (Baker, 1986). 5.1.2. Transmitter receptors in the AOB (Table 8) While many whole brain receptor localization studies include the MOB, the majority do not report results concerning the AOB. The most detailed studies find a strong signal for GABAA receptors in the glomerular, AEP, and AMC layers of the AOB (Zhang et al. 1991; Richards et al. 1987; Young and Kuhar, 1980b; Persohn et al. 1992; Laurie et al. 1992). For excitatory amino acid receptors, data exists for only NMDA, AMPA, and metabotropic receptors. A dense band of AMPA receptor subunits are found in the glomerular and external plexiform layers of the AOB (Petralia and Wenthold, 1992). NMDA receptor subunits are also present in the granule cell layer of the AOB (Petralia et al. 1994). Evidence also suggests the existence of metabotropic glutamate receptors in the AOB, but no laminar organization was given (Ohishi et al. 1993). There is not enough evidence to report the localization of modulatory neurotransmitter receptors in AOB. 538

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5.1.3. Outputs of the AOB The AOB has direct projections to the amygdala, specifically to the medial and posterior cortical nuclei, the bed nucleus of the stria terminalis and the nucleus of the accessory olfactory tract. These pathways may be involved in the processing of pheromonal information. Neurons in the AOB targets express gonadal steroid receptors and thus may be modulated directly by circulating hormones. The efferent connections of the accessory olfactory system are summarized in Fig. 22. 5.1.4. Centrifugal afferents to AOB There are major differences between centrifugal inputs to MOB and AOB. First, centrifugal inputs to AOB arise from far fewer brain regions than inputs to MOB. The major afferents to AOB are from the bed nucleus of the stria terminalis, the nucleus of the accessory olfactory tract, the medial amygdala nucleus and the posteromedial cortical amygdala nucleus (De Olmos et al. 1978; Shipley and Adamek, 1984). A restricted part of the medial division of AON sends a dense projection to the granule cell layer of AOB (Rizvi et al. 1992), but all other divisions of AON lack connections with AOB. 5.2. HIGHER ORDER CONNECTIONS OF THE ACCESSORY OLFACTORY SYSTEM AND REPRODUCTIVE FUNCTIONS Olfaction plays an important role in sexual behavior in many animals. Macrosomatic animals have a highly developed ability to use olfaction for identifying sexual partners, enemies and food; i.e., these animals use olfaction for survival and continuation of the species. The linkage between reproductive behavior and olfaction is not as strong in humans but we may still possess the neural hardware tying odors to sexual arousal and certainly the profit and loss statements of the fragrance industry attests to a key role of olfaction in human sex drives. In macrosomatic animals, the AOB is believed to be involved in processing pheromones that are initially transduced by vomeronasal neurons, which project to the AOB. The AOB projects to the anteromedial (MeAa) and posterior cortical (CoAp) nuclei of the amygdala, the bed nucleus of the stria terminalis and the bed nucleus of the accessory olfactory tract (De Olmos et al. 1978; Scalia and Winans, 1975; Shipley and Adamek, 1984) (Fig. 22). MeAa and CoAp project to other amygdaloid nuclei, notably the posterior nucleus of the amygdala (PA) (Cameras et al. 1992), and to the preoptic area and the hypothalamus (cf. Shiosaka et al. 1983 for review) (Fig. 22). The PA appears to receive convergent input from both the MeA and CoAp and projects heavily upon some of the same structures targeted by MeA and CoAp, namely the medial preoptic area and the ventromedial hypothalamic nucleus. Some of these secondary olfactory connections strongly influence sexual drive and the neurons involved in the connections contain steroid receptors and release peptides that mediate the sexual responses. For example, the posterodorsal part of MeA (MeApd) contains neurons that project to four cell groups that are known to be sexually dimorphic and differ in their roles in reproduction. The medial preoptic nucleus (MPO) is one of the sexually dimorphic targets of MeApd and lesions of MPO decreases male copulatory behavior (cf. Simerly et al. 1989, for review). Estrogen regulates the expression of CCK at the mRNA level in cells of MeApd; many cells containing cholecystokinin in MeApd project to MPO. In the female rat, CCK injection in the medial preoptic region enhances luteinizing hormone secretion 539

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Fig. 22. Some of the higher order connections of the accessory olfactory system. Emphasis is on possible circuits that mediate output responses such as autonomic or hormonal changes. Output projections of the AOB are shown as thick lines; higher order connections are shown as thin lines. Cortical structures are depicted as boxes; subcortical structures as ellipses. Abbreviations: AOB = accessory olfactory bulb; Me = medial amygdaloid nucleus; A O N m = anterior olfactory nucleus (m = medial division); BAOT = nucleus of the accessory olfactory tract; Bar = Barrington's nucleu; BST = bed nucleus of the stria terminalis; Ce = central nucleus of the amygdala; DB = nucleus of the diagonal band; D R = dorsal raphe nucleus; Hi = hippocamus; I M L = intermediolateral cell column of the thoracic level of the spinal cord; LC = locus coeruleus; M D = medial dorsal nucleus; M P O = medial preoptic area; M n R = median raphe; PAG = midbrain periaquaductal gray; PCo = posterior cortical amygdaloid nucleus; Po = posterior amygdaloid nucleus; Pit = pituitary; Pir = piriform cortex; P/Amb = nucleus ambiguus and periambiguual area; RVL = rostroventrolateral medulla; SCls = lumbrosacral spinal cord; SO = supraoptic nucleus; Tu = olfactory tubercle; TT = taenia tecta; V M A = ventromedial nucleus of the hypothalamus; 10 = dorsal motor nucleus of the vagus.

(Kimura et al. 1987) although its action on male sexual responses is not known. Nevertheless, this pathway could provide the neuroendocrine and anatomical substrates to regulate copulatory behavior in male rats. Recent research has utilized the cell activation marker properties of c-fos, an immediate-early gene, to demonstrate the specificity and pathways of the accessory olfactory system. For example, social odors activate cells in AOB (Schellinck et al. 1993) and, more specifically, central nuclei of the accessory olfactory system regulate species specific mating behavior (Fiber et al. 1993). The complexity of olfactory and somatosensory 540

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related integration has been shown in male rats after sexual interaction with estrous females whereby cells in the bed nucleus of the stria terminalis and medial preoptic area are activated (as visualized by c-fos expression) by inputs from the medial amygdala (probably via the accessory olfactory system) and the central tegmental field (probably via the genital somatosensory pathways) (Baum and Everitt, 1992). 5.3. SEXUAL DIMORPHISM OF AOB AND ITS TARGET STRUCTURES The growth of the AOB is influenced by gonadal steroids (Roos et al. 1988). The AOB of the male rat is significantly larger than that of females but if the male is castrated early in development, the AOB has a similar size to that of females. These findings correlate with sexual dimorphisms in other structures (e.g. preoptic area, ventromedial hypothalamic nucleus, medial nucleus of the amygdala, bed nucleus of the accessory olfactory tract and bed nucleus of the stria terminalis, medial region) that are known to influence sexual behavior (cf. Arnold and Gorski, 1984; Segovia and Guillam6n, 1993 for review), and receive direct or indirect projections from AOB (Scalia and Winans, 1975; Simerly et al. 1989).

6. 'NON-OLFACTORY' MODULATORY INPUTS TO THE OLFACTORY SYSTEM The olfactory system is heavily targeted by inputs from non-olfactory subcortical modulatory systems. These inputs arise from three sources; the nucleus of the diagonal band, dorsal and median raphe nuclei and locus coeruleus. The nucleus of the diagonal band (DB) is a component of the basal forebrain magnocellular system including DB, nucleus basalis and the medial septum. These basal forebrain neurons innervate most regions of the neocortex, the hippocampus and many other forebrain regions including the amygdala and the thalamus. The nucleus locus coeruleus and the dorsal and median raphe nuclei innervate cortical and subcortical structures throughout the CNS. 6.1. CHOLINERGIC INNERVATION OF THE OLFACTORY SYSTEM 6.1.1. Cholinergic inputs to the MOB

In the mouse about 3.5% of all neurons that project to the bulb originate in the horizontal limb of DB (Carson, 1984a); far fewer originate in the vertical limb of DB (Carson, 1984a; Shipley and Adamek, 1984) (Fig. 18 B,C). The vertical limb of the diagonal band is continuous with the horizontal limb; the horizontal limb is defined to begin where the diagonal band disappears from the surface and the nucleus is located deep to the olfactory tubercle (De Olmos et al. 1978). Many DB cells are cholinergic (Carson, 1984b; Macrides et al. 1981) but a double label study (Z~tborszky et al. 1986) showed that there are at least two distinct populations of DB neurons projecting to the olfactory bulb. About 20% of the DB neurons that project to the bulb are cholinergic; most of these cells are concentrated in the rostromedial portion of the horizontal limb of DB. At least as many DB bulbopetal neurons are GABAergic and are preferentially localized mainly in the lateral-caudal regions of the horizontal limb of DB (Z~borszky et al. 1986). Thus, there are at least two transmitter candidates in DB projections to the bulb. Accordingly, although the two types of neurons are somewhat segregated in DB, 541

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Fig. 23. Modulatory transmitter systems in the MOB. Darkfield photomicrographs showing the distribution of cholinergic (A), noradrenergic (B) and serotonergic (C) fibers to the layers of MOB. The axons are shown using antibodies to choline acetyltransferase (ChAT), dopamine-B-hydroxylase (DBH) and serotonin (5-HT) and immunocytochemistry.

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injections of anterograde tracers to label DB projections to the bulb (Luiten et al. 1987) may not distinguish the cholinergic from GABAergic input. Acetylcholinesterase (ACHE) is one marker for the cholinergic axons. AChE staining in the bulb preferentially concentrated in the IPL, GCL, inner third of the EPL, and the GL. Some glomeruli are more densely stained for AChE and correspond to regions of LHRH innervation (Zheng et al. 1988). The source of LHRH in these specialized glomeruli is unknown although Zheng et al. (1988) suggested vertical limb of DB as a possible source. The glomeruli containing dense AChE and LHRH label may include the modified glomerular complex as defined by Greer, Teicher and collaborators (Teicher et al. 1980; Greer et al. 1982). It has been suggested that these glomeruli may be areas of specialized olfactory processing during development (Teicher et al. 1980). AChE axon and terminal staining in MOB is partially confounded by the presence of AChE-positive cholinoceptive neurons in the bulb (Nickell and Shipley, 1988). A more suitable marker for cholinergic axons is choline acetyltransferase (CHAT), the requisite enzyme for acetylcholine synthesis. ChAT-stained axons are located in similar layers as described for AChE and are very fine in diameter (Fig. 23A) (Shipley et al. unpublished observations). The glomeruli of the AOB lack both ChAT and AChE staining. Thus, this represents another example where glomerular innervation differs in the AOB and MOB. The GABAergic projection from DB (Zaborszki et al. 1986) is more difficult to characterize than the cholinergic because the intrinsic GABAergic periglomerular and granule cells in the bulb provide such a massive intrinsic GABAergic innervation of the bulb. Of the HDB neurons projecting to the olfactory bulb, about 30% contain GAD, the synthetic enzyme for GABA (Rye et al 1984; Brashear et al. 1986; Zaborszky et al. 1986). The identified cholinergic and GABAergic neurons in HDB projecting to the bulb account for 40-50% of HDB neurons. Immunohistochemical procedures may not have labeled all ChAT or GAD containing cells; however, it is also possible that the remaining HDB neurons projecting to the bulb contain one or more other transmitters. Since there are intrinsic GABAergic neurons in all parts of the olfactory bulb (Macrides and Davis, 1983), it is difficult to determine whether the GABA fibers from HDB are distributed in the same way as the cholinergic input from this nucleus. Similarly, determination of the physiological role of the extrinsic GABAergic projection is complicated by the presence of intrinsic GABAergic neurons. Cholinergic receptors in M O B (Table 3) Regional patterns of cholinergic receptor localization in the MOB are in close agreement with terminal staining from the diagonal band of Broca. The internal and external plexiform layers show rather intense staining for ml, m3, and m 4 receptors (Buckley et al. 1988; Spencer et al. 1986; Fonseca et al. 1991; Rotter et al. 1979). Presynaptic m 2 receptors are immunocytochemically localized to periglomerular cells in the GL and tufted cells in the EPL (Fonseca et al. 1991). M2 receptors are also present in the IPL and granule cell layers. Nicotinic cholinergic receptors (nAChR) show a different regional distribution throughout the MOB. Autoradiographic and immunocytochemical studies report nAChR's to be located in the glomerular and external plexiform layers (Sahin et al. 1992; Hunt and Schmidt, 1978). Cholinergic actions in M O B There have been very few studies of cholinergic influences on the bulb. Electrical 543

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stimulation of the horizontal limb of the diagonal band (HDB) produces a biphasic field potential in the bulb consisting of an initial positive deflection followed by a slower negative wave. Repetitive stimulation of HDB neurons at 10 Hz causes marked potentiation of the negative portion of the response. Analysis of the depth profile of the field potential suggests that the negative wave (in GCL) results from excitatory currents flowing into granule cells. Consistent with this interpretation, mitral cells are inhibited during potentiation of the field potential (Nickell and Shipley, 1988). HDB neurons fire repetitively at 6-8 Hz (theta frequency) during investigative behavior which involves active sniffing. Thus it is possible that HDB cells provide some level of mitral cell inhibition during investigatory behavior. This might allow odor driven responses of mitral cells to predominate over spontaneous activity. More recently, HDB stimulation was also found to inhibit commissural fibers (Nickell and Shipley, 1993). Stimulation of HDB profoundly reduces the field potential caused by stimulation of the anterior commissure (AC). This appears to be due to presynaptic inhibition of AC terminals as during HDB stimulation, the threshold for antidromic activation of AC terminals increases dramatically. These effects are blocked by the muscarinic antagonist, scopolamine. These findings suggest that the cholinergic input from HDB to the bulb may function, in part, to regulate transmission of olfactory information between the two hemispheres. This is interesting in light of studies showing that AC fibers are required for access and recall of olfactory memories between the two hemispheres.

6.1.2. Cholinergic inputs to the piriform cortex Cholinergic inputs to PC arise from neurons in the horizontal limb of the nucleus of the diagonal band. Retrograde tracing studies demonstrate that diagonal band neurons that innervate PC are co-distributed among, but distinct from HDB neurons that project to MOB. This finding suggests that cholinergic inputs to MOB and PC originate from separate populations of HDB neurons. Due to specificity and sensitivity problems with antibodies raised against the synthetic enzyme for acetylcholine, choline acetyltransferase, it has been very difficult to render fine caliper cholinergic axons and terminals with immunohistochemical techniques. However, Lysakowski et al. (1989) achieved considerable success with immunohistochemical staining of fine cholinergic processes with ChAT and generated a comprehensive survey of cholinergic cortical innervation. This study reported that cholinergic inputs along the rostrocaudal axis of PC are fairly homogenous. Layer I receives a sparse cholinergic innervation that is considerable weaker than the corresponding innervation of other neo and paleocortical areas. Layer II and III, by contrast, receive a moderate and fairly uniform cholinergic input, although the density of fibers is heavier in layer II than in layer III. The pattern of cholinergic innervation of PC contrast with AChE staining in this structure, which is very heavy in layer I-III. Cholinergic receptors in the piriform cortex (Table 6)

The pattern of cholinergic receptor sites in PC is similar to the efferent inputs of HDB neurons. An array of anatomical techniques show all four muscarinic subtypes (m~, m2, m3, and m4) to be present in layer II PC (Buckley et al. 1988; Spencer et al. 1986; Zilles et al. 1989; and Rotter et al. 1979). The m~ receptor subtype has also been identified immunocytochemically to be on dendritic spines in layer I, but the identification of these 544

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dendrites were not determined (Levey et al. 1991). Nicotinic cholinergic receptors are located in layers II and III of PC, but as with the muscarinic receptors, the cellular and dendritic locations of these receptors are not known (Seguella et al. 1993; Hill et al. 1993; Sahin et al. 1992).

Cholinergic actions in the piriform cortex Hasselmo and Bower (1992a,b) have examined the actions of ACh on PC neurons excitability. In PC slices, ACh and muscarinic cholinergic agonists cause a suppression

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of intrinsic (layer Ib) fiber transmission without affecting transmission at afferent (layer Ia) fiber synapses. This suppression is presynaptically mediated by ACh actions on the M1 muscarinic subtype. Cholinergic agonists also appear to directly increase the excitability of pyramidal neurons, increasing neuronal bursts induced by intracellular depolarization. This effect is similar to that described for cholinergic agonists in the hippocampus and other cortical neurons, causing increased membrane excitability by blocking the slow afterhyperpolarization mediated by a calcium-dependent potassium current. 6.2. NORADRENERGIC (NE) INNERVATION OF THE OLFACTORY SYSTEM 6.2.1. NE innervation of the MOB

A significant modulatory input to the bulb is from the pontine nucleus, locus coeruleus (LC). In the rat all LC neurons contain the neurotransmitter, norepinephrine (NE); LC is the largest NE cell group in the brain. Shipley et al. (1985) estimated that up to 40% of LC neurons (400-600 of a total of 1,600 LC neurons) project to the bulb in the rat. The axons of neurons in LC project mainly to the infraglomerular layers of the bulb, particularly the internal plexiform and granule cell layers (McLean et al. 1989) (Fig. 23B). The external plexiform and mitral cell layers are moderately innervated while the glomerular layer is nearly devoid of NE input. This highly specific laminar innervation, unusual for LC terminal fields, pattern is observed in both MOB and AOB. In AOB, the internal plexiform layer is, in fact, sharply demarcated by the dense NE fibers running through it just deep to the multicellular output cell layer (mitral cell layer) (McLean et al. 1989) (Fig. 24). Based on these light microscopic studies it was suggested that the major target of the NE input is granule cells (McLean et al. 1989). NE receptors in MOB (Table 3)

Noradrenergic beta and alpha receptors have been identified anatomically in three layers of the MOB (Wanaka et al. 1989; Booze et al. 1989; Sargent-Jones et al. 1985; Nicholas et al. 1993; Young and Kuhar, 1980a). The granule cell layer, which receives a major noradrenergic input from LC, contains alpha2 and both beta subunits. When activated, these receptors are believed to alter the excitability of granule cells in the GCL. Both alpha receptor subtypes are localized to the external plexiform layer, while the glomerular layer and the IPL contain beta2 receptors. Physiological actions of N E in MOB

The nature of the physiological actions of NE in the bulb is a matter of controversy. In dissociated cultures of rat MOB neurons, NE has been reported to inhibit excitatory transmission from mitral cell to granule cells via presynaptic, alpha receptor mediated mechanisms (Trombley, 1992, Trombley and Shepherd, 1992). This mechanism appears to involve a decrease of calcium influx into the presynaptic terminal and involves a G-protein-coupled second messenger system and it was suggested that the alpha2 receptor is involved. In these same studies, NE was also found to decrease spontaneous GABAergic IPSPs in presumed mitral cells by an alpha receptor mediated mechanism. However, as will be discussed below, recent evidence indicates that alphal receptors are especially highly expressed in the intact MOB. Moreover, alpha2 receptors are generally 546

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Fig. 25. GABA neurons in piriform cortex (PC). Low (A) and high (B) magnification photomicrographs showing GABA-positive immunoreactive neurons in PC. Note that GABA-positive cells are distributed fairly uniformly throughout PC, except for the relatively low density of GABA-positive neurons in layer Ib. Section in B was processed for silver intensification of GABA immunoreactivity to highlight the morphological features of GABAergic interneurons.

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thought to be presynaptic. Thus the results of these experiments on dissociated olfactory bulb neurons are, at present, difficult to interpret. Salmoiraghi et al. (1964) and McLennan (1971) reported that iontophoretic applications of NE and NE agonists decreased mitral cell firing rates. This was interpreted to mean that NE causes granule cells to increase GABA release, thus inhibiting mitral cells. McLennan further showed that NE failed to decrease mitral cell firing when bicuculline, a specific antagonist of the GABAA receptor was co-applied with NE. However, using an in vitro whole turtle bulb preparation Jahr and Nicoll (1982) found that NE application caused increased mitral cell firing. Previous studies have measured the actions of exogenously applied NE. With exogenous application, issues of NE concentration, time of exposure and site of action are difficult to compare to the in vivo situation where NE is released at LC synapses in a pulsitile fashion that correlates with the firing rates of LC neurons. Therefore, we recently investigated the actions of NE on bulb neurons during physiologically confirmed activation of locus coeruleus. The experiments demonstrate that activation of LC causes a two-fold increase in the response of mitral cells to weak but not strong shocks applied to the olfactory nerve (Jiang et al. 1993). This is consistent with the idea that NE preferentially enhances responses to weak stimuli. Thus, when LC is activated by novel or unanticipated events, there may be a transient increase in mitral cell sensitivity to weak, odors. This could allow the animal to detect low level but potentially important odor cues, such as a predator or a pup straying from the nest. NE input to the AOB appears to have a very interesting function in mice. Bruce (1960) found that recently mated female mice will abort if presented with the odors of a strange male mouse that is not the mate. This effect is blocked if the NE input to the female's AOB is removed immediately after mating, presumably before olfactory memories of the mate are formed (Keverne and de la Riva, 1982; Rosser and Keverne, 1985). Thus, NE appears to be important in strengthening the memory of the odor of the 'husband'. The mechanism of memory formation has been examined by Kaba, Rosser and Keverne (1989) in the context of pregnancy block. They suggest that the dendro-dendritic synapse between granule cells and mitral cells in AOB may be critical for the memory formation and that NE by enhancing the inhibition of a subset of mitral cells for several hours following mating may facilitate a selective odor memory. They further suggest that, as a consequence of such neural activity, presenting the pregnant female with the stud male would produce activity of mitral cells matching that produced around the time of mating while strange males would produce different patterns of mitral cell activity leading to neuroendocrine responses that abort the pregnancy. NE has also been shown to be necessary for other olfactory memories such as maternal recognition in sheep (Pissonnier et al. 1985) and odor preference in young rats (Sullivan et al. 1989). Noradrenergic fibers arrive in the bulb before birth and increase in density (McLean and Shipley, 1991) at a time when olfactory bulb circuits are still being established in the bulb. The timing of noradrenergic axon arrival in MOB correlates with pharmacological evidence of noradrenergic influence on mitral cell excitability in the immature bulb (Wilson and Leon, 1988).

6.2.2. NE inputs to the piriform cortex Compared to the bulb, our knowledge of monoaminergic inputs to PC is limited. LC neurons project heavily to PC (Mason and Corcoran, 1979, Fallon and Loughlin, 1982) and LC lesions decrease NE levels in PC by 77% (Fallon and Moore, 1978). Like the 548

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projection to the neocortex, the LC projection to PC is primarily ipsilateral with a small contralateral component. Other brainstem and medullary noradrenergic cells groups do not appear to innervate PC. In the rat neocortex, NE fibers terminate with a degree of laminar organization, but with very little regional specificity; by contrast, NE fibers innervate the gyrencephalic neocortex of the primate with a higher degree of interlaminar and regional specificity (see Morrison et al. 1984 for review). Unfortunately, studies of similar detail in rat PC are not available. The primary study cited in this regard is the histofluorescence study of Fallon and Moore (1978) which could not unambiguously distinguish among 5-HT, DA and NE. Recently, we (Ennis et al. 1992) have investigated NE innervation of PC in considerable detail using immunohistochemical methods (Fig. 27). NE fibers in layer I were oriented in parallel to the pial surface. Similar to the noradrenergic innervation of parietal and sensorimotor cortices, layers Ia and III of PC contained a moderate plexus of NE fibers; layer II was sparsely innervated. A distinctive feature of NE innervation of PC is the long fibers oriented primarily parallel to the pial surface in layer Ia. The density and laminar distribution of NE fibers are relatively uniform along the rostrocaudal axis of PC. Adrenergic receptors in the piriform cortex (Table 6)

Both beta and alpha receptors are present in PC. In fact, both receptor types and their subunits are specifically localized in layer II of PC, while some studies see possible expression of the beta subtype in layer I (Nicholas et al. 1993; Wanaka et al. 1989; Sargent-Jones et al. 1985; Young and Kuhar, 1980a; Palacios and Kuhar, 1982; Unnerstall et al. 1984). This pattern of adrenergic receptor localization suggests that noradrenergic terminals from the nucleus locus coeruleus make synaptic connectivity in this layer of PC. Physiological actions of N E in the piriform cortex

The physiological action of NE on PC neurons has received relatively little attention by comparison to numerous studies in other cortical regions. Sheldon and Aghajanian reported that NE caused excitation of putative interneurons located at the layer II/III border (Sheldon and Aghajanian, 1990). NE has also been shown to block the slow afterhyperpolarization in guinea-pig pyramidal cells (Constanti and Sim, 1987), an effect similar to NE actions on hippocampal pyramidal neurons. 6.3. SEROTONIN (5-HT) INNERVATION OF THE OLFACTORY SYSTEM 6.3.1. 5-HT innervation of the MOB

The midbrain dorsal and median raphe provide strong inputs to the main olfactory bulb. In the rat, about 1,000 dorsal and 300 median raphe neurons innervate MOB. These neurons are serotonergic and do not contain tyrosine hydroxylase (McLean and Shipley, 1987b) or substance P (Magoul et al. 1988). The raphe input enters MOB via the olfactory peduncle and perhaps via the olfactory nerve layer (McLean and Shipley, 1987b). Thick serotonergic fibers preferentially innervate the glomeruli of MOB while thinner serotonergic axons preferentially innervate inframitral layers (McLean and Shipley, 1987b) (Fig. 23C). In neocortex, thin axons originate in the dorsal raphe and

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thick axons from median raphe neurons (Mamounas and Molliver, 1988) so one may assume the same type of arrangement occurs in MOB. Serotonergic axons do not innervate the glomeruli of AOB (Fig. 24)just as cholinergic axons avoid this layer. Since the AOB has far fewer PG cells than in MOB, the paucity of 5-HT and ACh innervation of the AOB glomerular layer suggests that serotonergic and cholinergic inputs target PG cells in MOB; the relative absence of PG neurons in AOB, thus, might account for the lack of serotonergic/cholinergic input to that layer in AOB. Interestingly, the serotonergic innervation of MOB glomeruli in primates is considerably reduced (Azmitia and Gannon, 1986; Takeuchi et al. 1982) compared to other mammals that have been studied such as rats (McLean and Shipley, 1987b), cats (Takeuchi et al. 1982), rabbits (personal observation) and hamster (Kream et al. 1984). Whether this reflects a true species difference or differences in technique is unknown. Recently, it shown that 5HT is necessary for the acquisition and expression of conditioned olfactory learning in neonatal rats (McLean et al. 1993). 5 - H T receptors in the M O B (Table 3)

The cells upon which serotonergic axons synapse in the olfactory bulb are becoming elucidated. Most of the earlier information is based on location of receptor binding which may lead one to infer that certain cell types receive serotonergic input. More recently, the cells which express mRNA for serotonergic receptor subtypes have been identified which has helped identify the cells receiving serotonergic input even further. In the case of the olfactory bulb, there are mainly two subtypes present; 5-HT1A and 5-HT2A. The receptors for the 5-HTIA subtype are located in the external plexiform layer, mitral cell layer and minimally in the granule cell layer while the cells expressing the mRNA for the 5-HTIA receptor are located in the mitral and granule cell layers (Pompeiano et al. 1992). This has lead to the speculation that cells in the mitral and granule cell layers possess the 5-HT~A receptor. It is possible that many of the cells possessing the receptor are glia since a recent immunocytochemistry study has shown that the 5-HT1A receptor is localized to glial cells (Whitaker-Azmitia et al. 1993). The 5-HT2A receptor is found mainly in the external plexiform layer. On the other hand, the mRNA for the 5-HT2A receptor has been shown by in situ hybridization to be in the mitral cell and external plexiform layers (Pompeiano et al. 1994) and more precisely, in mitral and tufted cells (McLean et al. 1994). This leads one to speculate that the dendrites or cell bodies of olfactory bulb output cells receive serotonergic input via 5-HTzA receptors. It is interesting that 5-HTIA and 5-HTzA receptor binding are both relatively low in the glomerular layer where there is substantial 5-HT innervation and possibly, synaptic input onto output cells (Liu and Shipley, in preparation). This could mean that a 5-HT receptor subtype other than the 5-HT~A or 5-HT2A is responsible for receiving 5-HT input in the glomeruli or it could mean there is a receptor/axon mismatch for 5-HT input in the bulb. Another 5-HT receptor subtype, 5-HTzc is in low quantities in the olfactory bulb. The accessory olfactory bulb has not been analyzed adequately for the 5-HT receptor subtypes but it is clear that the mRNA for the 5-HT2A receptor is low to negligible in AOB (McLean et al. 1994). 6.3.2. 5-HT inputs to the piriform cortex

As with the other monoamines, a comprehensive study of 5-HT inputs to PC has not been undertaken. Anterograde and retrograde tracing studies have demonstrated a rich 551

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projection from the dorsal raphe nucleus to PC (De Olmos and Heimer, 1980; Vertes, 1991). Ascending serotonergic fibers from the dorsal raphe terminating in PC are believed to travel in the ventrolateral aspect of the medial forebrain bundle (Azmitia and Segal, 1978). Anterograde labeling of ascending dorsal raphe axons demonstrated that the entire PC was targeted by raphe projections (Vertes, 1991). The projection was reported to be heavier to rostral than caudal PC, and heavier to the deeper than the superficial layers. However, the transmitter nature of labeled fibers were not be identified in this anterograde tracing study. Our recent immunocytochemical studies indicate that the 5-HT innervation of the cortex exhibits less interlaminar specificity than NE terminals and is, in general, complimentary to that of NE inputs (Fig. 27). The serotonergic innervation of PC is very heavy by comparison to DA and NE. 5-HT fibers are especially heavy in layers I and III and the endopiriform nucleus. The density of 5-HT fibers progressively decreases in the deeper parts of layer III. Layer II by contrast, is sparsely innervated. 5-HT fibers in all layers are relatively short, with a tortuous, convoluted orientation and exhibit more varicosities than NE or DA fibers. As with NE, there is little variation of 5-HT fiber distribution or density at different rostrocaudal levels of PC. 5 - H T receptors in the piriform cortex (Table 6)

Less information is available concerning the 5-HT receptor subtypes present in parts of the olfactory system other than in the olfactory bulb. However, it is clear that the 5-HT2c receptor subtype is very densely distributed in the anterior olfactory nucleus as revealed by receptor binding (Mengod et al. 1990a) and in situ hybridization (Mengod et al. 1990a; Hoffman and Mezey, 1989) while the 5-HTzAreceptor subtype is especially dense in the external division of the anterior olfactory nucleus (Mengod et al. 1990b; Pompeiano et al. 1994; McLean et al. 1994) and present in moderate density in other divisions of AON (Mengod et al. 1990b; Pompeiano et al. 1994). The 5-HTIA receptor binding is similar in distribution to that of the 5-HT2A receptor in the AON with a special note that the mRNA for 5-HT~A is in extremely high concentration in external division, just as the 5-HTzA receptor is. Results from binding and in situ hybridization studies in the piriform cortex suggest that 5-HT1A receptors are located on the dendrites of pyramidal cells (layer II) and also on intrinsic cells of layer III (Pompeiano et al. 1992). In the entorhinal cortex, the 5-HT~A receptors have been suggested to be on stellate cells in layer II and on granule cells in layers V and VI (Pompeiano et al. 1992). Regarding the 5-HTzA and 5-HTzc receptors, in situ hybridization revealed strong signal of both receptors over cells in layer II and III (Pompeiano et al. 1994). Physiological actions of 5-HT in the piriform cortex

The actions of 5-HT in PC have recently been examined by Aghajanian and colleagues (Sheldon and Aghajanian, 1990, 1991; Gellman and Aghajanian, 1994). 5-HT has several actions on PC pyramidal cells as well as putative interneurons that are mediated by distinct 5-HT receptor subtypes. 5-HT has mixed, but predominately excitatory actions on layer II pyramidal cells. This excitation appears to be mediated by a reduction of the so called M-current, a non-inactivating voltage-dependent outward potassium current, and is mediated via 5-HTlc receptors. 5-HT also causes inhibition and an increase in the frequency of IPSPs in other pyramidal cells. This is caused by excitation 552

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of putative GABAergic interneurons that are directly activated by 5-HT acting at the 5-HT2 receptor subtype. 6.4. DOPAMINE (DA) INNERVATION OF THE OLFACTORY SYSTEM There is no known extrinsic DA innervation of the olfactory bulb. As noted above, however, the MOB contains several hundred thousand intrinsic juxtaglomerular DA neurons

6.4.1. Dopamine (DA) innervation of the piriform cortex The dopaminergic innervation of PC exhibits a marked rostrocaudal gradient and laminar specificity (Fig. 27). Rostrally, DA fibers are relatively sparse and primarily confined to layer III. Along the rostral to caudal axis of PC, the density of innervation progressively increases and DA fibers invade more superficial layers of PC. By the caudal limits of PC, a moderately dense plexus of DA fibers extends from the deep part of layer I through layer III. DA receptors in the pirijorm cortex

While the source of dopaminergic input to PC is unclear, in situ and immunocytochemical studies have found the presence of D1 receptors in layer II of PC (Mansour et al. 1990; Fremeau et al. 1991; Huang et al. 1992). Little is known, though, about the significance of these receptors to PC physiology. To date, there is little definitive evidence concerning the presence of D3 receptors in PC. 6.5. COMPARISON OF NE, 5-HT AND DA INPUTS IN THE RAT P I R I F O R M CORTEX (Fig. 27) The monoaminergic innervation of the cerebral cortex is much denser than was appreciated in the 1970s and early 1980s. We have initiated an immunocytochemical study of NE, 5-HT and DA innervation of PC. Our preliminary findings show that all three monoamines robustly innervate the entire rostrocaudal extent of PC. Of particular note was the surprisingly dense terminal fields of these monoamines in PC which were equal to, and in many cases, greater than their corresponding innervation of medial prefrontal, sensorimotor, parietal, insular and hippocampal cortices. As with other cortical regions, 5-HT provided the densest innervation of PC, followed by NE and then TH (5-HT > NE > DA). NE fibers in PC show some laminar preferentially but do not have the same degree of laminar specificity as in the bulb. 6.6. D I F F E R E N T I A L INNERVATION OF MOB AND AOB Some centrifugal afferents are common to both MOB and AOB. These include the subcortical modulatory systems: diagonal band, dorsal and median raphe nuclei and locus coeruleus. The terminal distribution of these common inputs differ in AOB and MOB, especially with respect to the cholinergic and serotonergic inputs. In AOB, both the cholinergic and serotonergic inputs avoid the glomeruli whereas they heavily innervate the glomeruli of MOB. The cholinergic-serotonergic inputs to AOB are mainly to the granule cell layer and internal plexiform layer (McLean and Shiplcy, 1987b; Le Jeune 553

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and Jourdan, 1991). The NE input appears to have similar laminar termination patterns in both MOB and AOB.

7. A C K N O W L E D G M E N T S

Supported by grants from NIH NIDCD DC00347, DC02588 NINDS NS24698, NS29218 and DOD DAMD17-91-C-1071 (M.T.S. and M.E.) and from M.R.C. of Canada #MT- 10931 (J.H.M).

8. ABBREVIATIONS ac

Acb AHC AI AOB AGL AGCL AMCL AONd AONe AON1 AONm AONvp BLA BLAp BMA BMAp CA1 CA3 CALB CeA CoAA CoApl CoApm CPu DG DP En Ent Entl Entre EPL GL GCL HDB ICj

anterior commissure nucleus accumbens anterior hippocampal continuation anterior insular cortex accessory olfactory bulb accessory olfactory bulb, glomerular layer accessory olfactory bulb, granule cell layer accessory olfactory bulb, mitral cell layer anterior olfactory nucleus, dorsal anterior olfactory nucleus, external anterior olfactory nucleus, lateral anterior olfactory nucleus, medial anterior olfactory nucleus, ventroposterior basolateral nucleus of the amygdala basolateral nucleus of the amygdala, posterior part basomedial nucleus of the amygdala basomedial nucleus of the amygdala, posterior part CA1 region of the hippocampus CA3 region of the hippocampus Calbindin central nucleus of the amygdala anterior cortical nucleus of the amygdala cortical nucleus of amygdala, posterolateral cortical nucleus of amygdala, posteromedial caudate/putamen dentate gyrus dorsal peduncular cortex endopiriform nucleus entorhinal cortex lateral division of the entorhinal cortex medial division of the entorhinal cortex external plexiform layer glomerular layer granule cell layer horizontal limb of the diagonal band islands of Calleja 555

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IL IPL LA LH lot MCL MeAa MeApd MeApv NLOT ONL opt PA PAA Pir PLCo PMCo Pre RF SEZ SO st

Sub Tr TTd TTis TTvs Tu VDB VN VP

infralimbic cortex internal plexiform layer lateral nucleus of the amygdala lateral hypothalamic area lateral olfactory tract mitral cell layer medial nucleus of the amygdala, anterior part medial nucleus of the amygdala, posterodorsal part medial nucleus of the amygdala, posteroventral part nucleus of the lateral olfactory tract olfactory nerve layer optic tract posterior nucleus of the amygdala piriform-amygdala area piriform cortex posterolateral cortical amygdala nucleus posteromedial cortical amygdala nucleus presubiculum rhinal fissure subependymal zone supraoptic nucleus stria terminalis subiculum transitional area taenia tecta, dorsal taenia tecta, inferior-superior taenia tecta, ventral-superior olfactory tubercle ventral limb of the diagonal band vomeronasal nerve ventral pallidum

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Subject index

accessory olfactory bulb architecture 536 external plexiform layer 537 glomerular layer 537 internal plexiform layer 537 mitral cell layer 537 neurotransmitters 537 outputs 539 periglomerular cells 537 transmitter receptors 538 acetylcholine cerebellum 113, 127, 163 inferior olive 242 nucleus of the diagonal band neurons 541 striatal neurons 394 acetylcholinesterase basal interstitial nucleus 163 juxtaglomerular cells 486 Golgi cells 88, 118, 127 parasagittal zones 195 Purkinje cells 23, 127 adenosine Purkinje cells 77, 79 adenosine deaminase cerebellum 78 adenosine kinase cerebellum 78 adenosine receptors al in cerebellum 78 a2 in striatum 449 adrenergic receptors in anterior olfactory nucleus 514 in cerebellum 107 in olfactory bulb 546 aldolase C in Purkinje cells 44 allocortex inputs to striatal patch/

matrix 438 olfactory cortex 474 Alzheimer's disease olfactory deficits 473 amphetamine striatal dopamine receptors 449 anterior hippocampal continuation 516 anterior lobe, cerebellum 135 anterior olfactory nucleus architecture 509 functions 515 inputs 509 outputs 509 pars externa topography 510 projection to hypothalamus 528 transmitter receptors 514 transmitters 514 tufted cell input 491 aspartate accessory olfactory bulb 537 anterior olfactory nucleus 514 cerebellar nuclei 160 climbing fibers 55 juxtaglomerular cells 486

basal ganglia diseases 371 inputs, cortico-cortical organization 413 organization 372, 409, 410 basal interstitial nucleus (Langer) 143, 146, 163 basal pontine gray collaterals to cerebellar nuclei 302 mossy fibers 291,301 bed nucleus of the stria termi-

nalis accessory olfactory input 539 benzodiazepine receptors cerebellum 94 Bergmann glia s e e cerebellar cortex Blanes cells 491 Bowman's glands 470 brachium conjunctivum 284 breast cancer cerebellum 48 butyrylcholinesterase Bergmann glial cells 130 Ca2+-ATPase in Purkinje cells 31 calbindin anterior olfactory nucleus 514 juxtaglomerular cells 486 Purkinje cells 36 calcitonin gene-related peptide (CGRP) climbing fibers 275 mossy fibers 305 Purkinje cells 23 calmodulin in Purkinje cells 36, 37 calreticulin in Purkinje cells 32 calretinin anterior olfactory nucleus 514 climbing fibers 283 juxtaglomerular cells 486 mitral cells 490 somesthetic corpus cerebelli 207 stellate/basket cells 85 unipolar brush cells 90 calsequestrin in Purkinje cells 31

575

Subject index cannabinoid receptor

in striatum 450 carbonic anhydrase

oligodendrocytes 130 carbon monoxide

cerebellum 77 carnosine

olfactory receptor neurons 475 central amygdalar nucleus

input from insular cortex 534 central cervical nucleus

collaterals to cerebellar nuclei 302 spinocerebellar pathway 287, 289 cerebellar commissure 284 cerebellar cortex

astroglia 128, 130 basket (stellate) cell appearance 15 calretinin 85 cyclic GMP 77 immunoreactivities 84 nitric oxide synthase 77 parvalbumin 85 Bergmann glia AMPA receptors 62 butyrylcholinesterase 130 cyclic GMP 77 FAL-epitope 41 3-fucosyl-acetyl-lactosamine (FAL) 132 GABAA receptors 100 glial fibrillary acidic protein 130 glutamate transporter 21 glycan phosphorylase 41 glycogen phosphorylase 41 guanylate cyclase 77, 133 NADPH-diaphorase 77 nitric oxide synthase 77, 133 5'-nucleotidase 130, 132 protein kinases 132 vimentin 130 zebrin I 131 zonal expression patterns 132 candelabrium cell

576

appearance 16 cholinergic input 113 f f climbing fibers see climbing fibers

columnar modules 301 connections 170 ff. corticonuclear projection and compartments, monkey 184, 188 and cytochemical zones, cat 177 and zebrins 203 corticovestibular projection 188 cytology 1 development 217 f f folial pattern 135 ff. fractured somatotopy 301 Golgi cells acetylcholinesterase 88, 118, 127 appearance 14, 85 choline acetyltransferase 89, 119 displaced 127 enkephalin 89 GABA 87 glycine 87 guanylate cyclase 77 metabotropic glutamate receptors 73 somatostatin 89 granule: cells appearance 4 GABAA receptors 93, 97 gephyrin 101 glutamate 58 guanylate cyclase 77 heme oxygenase-2 77 kainate receptors 61, 69 location 4 metabotropic glutamate receptors 75 migration and NMDA receptor 71 NADPH-diaphorase 77 nitric oxide synthase 77 NMDA receptors 61, 70 gross anatomy 133 ff. hypothalamic input (histamine) 110 interneurons 14, 81 lobes 133 ff.

longitudinal zones see cerebellar cortex, parasagittal zones Lugaro cell appearance 16, 89 GABA 89 medullary core 175 micro zone 176, 248, 249, 301 monoaminergic inputs dopamine 110 noradrenalin 105 serotonin 102 mossy fibers see mossy fibers

neuroglia 128 oligodendroglia carbonic anhydrase 130 cerebellar 128 output modular organization 176 parallel fibers adenosine receptors 78 arrangement 5 benzodiazepine receptors 94 GABAB receptors 101 glutamate 58 5'-nucleotidase 79 synaptology 4 parasagittal zones acetylcholinesterase 195 cyclic GMP-dependent protein kinase 219 cytochrome oxidase 189, 193 development 218 existence 170 3-fucosyl-acetyl-lactosamine (FAL) 193 GABA binding 100 HNK-1 antigen 189 motilin 190 nerve growth factor receptor 189 5'-nucleotidase 189, 191 parvalbumin 220 P-path antigen 189 protein kinase C 193 Purkinje cell specific glycoprotein 220 rat 201

Subject index

taurine 190 transmitter receptors 189 zebrins 189, 201 Purkinje cells acetylcholine effect 127 acetylcholinesterase 23 adenosine 77, 79 adenosine deaminase 78 adenosine kinase 78 adrenergic receptors 110 aldolase C 44 AMPA receptors 60 axon 9, 11, 175 axon collaterals 15 benzodiazepine receptors 94 Ca2+-ATPase 31 calbindin 36 calcitonin gene-related peptide (CGRP) 23 calmodulin 36, 37 calreticulin 32 calsequestrin 31 carbon monoxide 77 cerebellins 39 climbing fiber input see climbing fiber cyclic GMP-dependent protein kinase 34 cytochrome oxidase 38 dendritic tree 7, 13 diacylglycerol 24 enolase 43 GABA as transmitter 17 GABAB receptors 101 gangliosides 41 gephyrin 101 guanylate cyclase 34, 36, 77 heme oxygenase-2 77 histamine receptors 112 inositol- 1,4,5-triphosphate receptor 24, 29 L-7 39 lower vertebrates 13, 14 metabotropic glutamate receptors 73 monoclonal antibodies 41 motilin 21 nerve growth factor 44

nerve growth factor (NGF) receptors 41 NMDA receptor subunits 70 5'-nucleotidase 79 orphan glutamate receptors 72 P400 protein 27 paraneoplastic diseases 47 parvalbumin 36, 37 PEP-19 polypeptide 38 phospholipase C 25, 32 protein kinase C 32, 33 serotonin receptors 104 somatostatin 24 taurine 21 T-cell antigens 39 tyrosine hydroxylase 24 visinine 39 zebrins 39, 41 stellate cells appearance 15 calretinin 85 cyclic GMP 77 guanylate cyclase 77 immunoreactivities 84 parvalbumin 85 unipolar brush cell appearance 16, 89 calretinin 90 cholinergic input 117 glutamate 89 glutamate receptors 65 secretogranin 89 cerebellar nuclei afferents climbing/mossy fiber collaterals 165 ff. monoaminergic 167 peptidergic 168 Purkinje cell 164 aspartate 160 cat 146 caudomedial group 141 cerebellar input 138 cholinergic input 117 cholinergic neurons 121, 163 climbing fiber input 139 dentate nucleus see cerebellar.nuclei, lateral nucleus fastigial nucleus

cells 140 fastigio-bulbar tract 143 somatostatin 164 uncinate tract 143 globose nucleus 171 glutamate 160 glutamate receptors 60 ff. in lower vertebrates 14 interneurons (intrinsic) existence 159 glycine 160 interposed nucleus cells 140 cholecystokinin 164 lateral (dentate) nucleus cells 140 monoaminergic input 138 mossy fiber input 138 NADPH-diaphorase 77 nucleo-cortical projection as collaterals 158 different cell types of origin 159 glutamate 158 nucleo-olivary projection GABA 140, 154, 234 glycine 157 parvicellular regions 143 primates 148 projections 141 ff. rat 151 red nucleus input to 303 serotonergic input 140 serotonin receptors 104 subdivision 140 cerebellar projections and basal ganglia 375 cerebellins in Purkinje cells 39 cerebellum see cerebellar cortex, nuclei cerebral cortex input to basal ganglia 425 laminar input to striatum 437 c-fos striatal dopamine receptor drugs 447 chemosensory transduction olfactory system 470 cholecystokinin input to cerebellar nuclei 168

577

Subject

index

interposed nucleus cells 164 juxtaglomerular cells 486 mossy fibers 305 choline acetyltransferase cerebellum 113 Golgi cells 89, 119 mossy fibers 303 transient Purkinje cell expression 23 cholinergic receptors see nicotinic, muscarinic cilia olfactory 470 Clarke's column dorsal spinocerebellar tract 287 climbing fibers arrangement 11 aspartate 55 calcium-binding proteins 275 calretinin 283 collaterals to cerebellar nuclei 165 glutamate 55 homocysteate 57 inferior olive origin 12 longitudinal strips 243 parvalbumin 269 peptides 275 cocaine striatal dopamine receptors 449 cognition and odors 532 columnar modules, cerebellum 301 copula pyramidis, cerebellum 136 copulatory behavior 539 cortical amygdalar nucleus accessory olfactory input 539 corticopontocerebellar projection 291 corticotropin-releasing factor (hormone) anterior olfactory nucleus 514 climbing fibers 275 inferior olive 277

578

input to cerebellar nuclei 168 mitral cells 490 mossy fibers 286, 304 cuneate (internal) nucleus cuneocerebellar pathway 287 GABA input to inferior olive 234 cuneocerebeilar tract 287, 287, 289, 293, 295 cyclic GMP cerebellum 77 cyclic GMP-dependent protein kinase in Purkinje cells 34 cerebellar development 219 cytochrome oxidase parasagittal zones 189, 193

enkephalin climbing fibers 275 D2 receptor modulation, in striatum 447 Golgi cells 89 input to cerebellar nuclei 168 mossy fibers 304 enolase in cerebellum 43 entopeduncular nucleus (internal globus pallidus, primates) nomenclature 376 entorhinal cortex and olfactory memory 536 external cuneate nucleus cuneocerebellar projection 287

dentate nucleus see cerebellar nuclei, lateral nucleus diacylglycerol (DAG) in Purkinje cells 24 dopamine input to cerebellum 110 input to piriform cortex 553 juxtaglomerular neurons 481 dopamine receptors D 1 in piriform cortex 553 D1 in striatum 447 D1, D2 in cerebellum 110 D2 in main olfactory bulb D2 in striatum 447 dorsal accessory olive see inferior olive dorsal cochlear nucleus 143 dorsal raphe input to olfactory bulb 550 input to piriform cortex 552 dorsal striatum (caudate, putamen) striatal regional differences 456 dorsomedial nucleus (thalamus) see mediodorsal nucleus

FAL-epitope in Bergmann glia 41 fastigial nucleus see cerebellar nuclei fastigio-bulbar tract 143 floccular peduncle 146, 209 flocculus, cerebellum connections 209 location 136 food and olfactory system 539 fractured somatotopy, cerebellum 301 3-fucosyl-acetyMactosamine (FAL) Bergmann glia 132 parasagittal zones 193

enemies olfactory detection 539

GABA (gamma-aminobutyric acid) accessory olfactory granule cells 537 cuneate nucleus 234 Golgi cells 87 inferior olive inputs 231, 234 inferior olive interneurons 233 juxtaglomerular cells 481 Lugaro cells 89 nucleo-olivary projection 140, 154, 234 nucleus of the diagonal

S u b j e c t index band cells 541,543 nucleus prepositus hypoglossi 237 olfactory granule cells 492 parasolitary nucleus 234, 237 periglomerular cells 537 Purkinje cells 17 striatal 372 vestibulo-olivary projection 157, 234, 237 GABA receptors accessory olfactory bulb 538 anterior olfactory nucleus 515 cerebellum 93 ff. main olfactory bulb 493, 500 piriform cortex 529 striatal 451 GABA transporters in Bergmann glia 21 GAD (glutamic acid deear-

boxylase) Purkinje cells 17 striatal projection neurons 444 two forms, Purkinje cells 17

gangliosides in cerebellum 41

gephyrin cerebellum 101

glial fibrillary acidic protein cerebellar astroglia 130

globus pallidus, external segment (globus pallidus, nonprimates) aspinous neurons 396 output 399 projections 396 spiny neurons 396 striatal inputs 397 subthalamic nucleus 397

glohus pallidus, internal segment see also entopeduncular nucleus dual output to thalamus 424 location 376 glomerulus see specific structure

glutamate cerebellar nuclei 160 cerebellum 50 climbing fibers 55 corticostriatal 372 granule cell/parallel fiber 58 mitral cells 490, 507 nucleo-cortical projection 158 olfactory receptor neurons 475 piriform cortex pyramids 525

glutamate receptors anterior olfactory nucleus 515 cerebellum 60 ff. main olfactory bulb 493 piriform cortex 529 striatal 451

glutamate transporter cerebellum 50 mossy fibers 52

glutaminase

cerebellum 77, 133 in Purkinje cells 34, 36, 77

heme oxygenase-2 cerebellum 77

hemisphere, cerebellum 135 ff. histamine input to cerebellum 111

histamine receptors cerebellum 112

HNK-1 antigen parasagittal zones 189

Hodgkin's disease cerebellum 47

homocysteate climbing fibers 57

Huntington's disease basal ganglia 371,453

6-hydroxydopamine lesions striatal dopamine receptors 448

hypothalamus histaminergic input to cerebellum 111

mossy fibers 52

glutamine mossy fibers 52

glycan phosphorylase in Bergmann glia 41

glycine Golgi cells 87 interneurons, cerebellar nuclei 160 nucleo-olivary projection 157

glycine receptor cerebellum 101

glycogen cerebellar astroglia 130

glycogen phosphorylase in Bergmann glia 41

gracile (internal) nucleus cuneocerebellar pathway 287 granule cell see specific structure

group Y (Brodal) location 143 organization 209 projection to oculomotor nucleus 145

guanylate cyclase

immediate early genes striatal dopamine receptor drugs 449

induseum griseum connections 516

inferior olive afferents 233 axon terminals 231 calcium-binding proteins 275 catecholamine input 239 cholinergic input 242 climbing fiber origin 12 cuneate nucleus input 234 dorsal accessory subnucleus 172, 225, 228 electrotonic coupling 228 GABA interneurons 233 GABA terminals 231 medial accessory subnucleus 225 muscarinic receptors 242 nicotinic receptors 242 nucleo-olivary GABA projection 234 nucleo-vestibular GABA projection 234

579

Subject index

olivocerebellar projection chicken 244 longitudinal strips 243 mammals 225 parvalbumin 269 rat 256, 257 to anterior lobe 244 to caudal vermis, flocculus 265 to hemisphere 259 transmitters 275 ff. turtle parasolitary nucleus input 234, 237 parvalbumin 282 peptides 275 ff. principal subnucleus 225 raphe input 241 red nucleus input 261 serotoninergic input 231, 239 structure 225, 228 vestibular input 157 infralimbic cortex connections 518 inositol-l,4,5-triphosphate (IP3) receptor in Purkinje cells 24, 29 insular cortex olfactory input 528, 534 projection to central amygdalar nucleus 534 interposed nucleus see cerebellar nuclei intralaminar nuclei (thalamus) input to striatum 435 juxtaglomerular neurons types 476 transmitters 481

L-7 in Purkinje cells 39 lateral habenula entopeduncular input 430 lateral nucleus see cerebellar nuclei lateral reticular nucleus collaterals to cerebellar nuclei 302 mossy fibers 165, 291 lateral vestibular nucleus

580

cerebellar input 138 zonal arrangement of mossy fibers 293 limbic system input to striatal patch/ matrix 434, 437 lobulus simplex, cerebellum 135 locus coeruleus input to cerebellum 106 input to olfactory bulb 546 input to piriform cortex 550 long term depression (LTD) cerebellum 36, 73 macrosomatic mammals two olfactory systems 473 main olfactory bulb centrifugal inputs 507 cholinergic inputs 541,543 external plexiform layer components 486 tufted cells 486 Van Gehuchten cells 487 functional organization 496 glomerular layer components 475 excitatory transmitters 501 external tufted cell dendrites 480 functional units 496 glomerular inhibition 499, 500, 503 glomerulus core and shell 475 infraglomerular inhibition 499, 503 juxtaglomerular neurons 476, 499 local circuits 499 olfactory nerve dopamine modulation 494 summation of inputs 498 granule cell layer Blanes cells 491 granule cells 491 organization 491 transmitters 492 internal plexiform layer organization 490

intrabulbar association system 487, 510 mitral cell layer intrabulbar collaterals 504 organization 488 projection to piriform cortex 491 response to odors 490 transmitters 490 mitral-granule cell interactions 492 lateral inhibition 501 oscillations 502 self-inhibition 50 noradrenergic actions 546 noradrenergic inputs 546 olfactory nerve layer (ONL) carnosine 475 components 474 glutamate 475 olfactory marker protein 475 projections intrabulbar collaterals 504 to insular and perirhinal cortex 528 serotonergic inputs 551 subependymal zone 493 topography of odor properties 496 transmitter receptors cholinergic 543 excitatory amino acid 493 GABA 493 noradrenergic 546 serotonin 551 maternal recognition 548 medial amygdalar nucleus accessory olfactory input 539 medial frontal cortex see also infralimbic, prelimbic cortex olfactory input 534 medial preoptic nucleus male copulatory behavior 539 medial vestibular nucleus cholinergic mossy fibers 115, 304

Subject index

median raphe (superior central nucleus) input to olfactory bulb 550 mediodorsal nucleus (thalamus) olfactory input 529, 533 memory olfactory-related 536 olfactory and norepinephrine 548 microvillar cells olfactory epithelium 470 micro zones see cerebellar cortex middle cerebellar peduncle 284, 291 mossy fiber acetylcholine 114, 303 appearance 4, 6, 7 calcitonin gene-related peptide 305 cholecystokinin 305 choline acetyltransferase 303 collaterals to cerebellar nuclei 165, 302 concentric arrangement 284 corticotropin-releasing hormone 286, 304, 305 course 284 enkephalin 304 fractured somatotopy 301 glutamate 51,303 glutamate transporter 52 glutaminase 52 glutamine 52 origins 165, 285 ff. patchy mosaic 301 rosettes 4, 7, 303 semicircular fibers 284 serotonin 303 somatotopic organization 299 vestibulocerebellar 285, 304 zonal arrangement 293 motilin in Purkinje cells 21 parasagittal zones 190 motor activity and olfaction 534 MPTP

nigral lesions 453 muscarinic receptors anterior olfactory nucleus 514 cerebellum 123 inferior olive 242 main olfactory bulb 543 piriform cortex 544 striatum 450 subtypes 450 NADPH-diaphorase cerebellum 77 juxtaglomerular cells 481 neocortex input to striatal patch/ matrix 438 nerve growth factor (NGF) in Purkinje cells 44 nerve growth factor (NGF) receptors in Purkinje cells 41 parasagittal zones 189 neuropeptide-Y juxtaglomerular neurons 481 neurons in olfactory granule cell layer 492 neurotensin accessory olfactory bulb 537 nicotinic receptors anterior olfactory nucleus 514 cerebellum 121 inferior olive 242 main olfactory bulb 543 piriform cortex nigrostriatal projection dopamine 376 nitric oxide (NO) and Purkinje cells 36 cerebellum 76 nitric oxide synthase (NOS) in cerebellum 36, 76, 133 noradrenaline input to cerebellum 105, 167 nucleo-cortical projection 158 nucleo-olivary projection 154 5'-nucleotidase cerebellum 79 parasagittal zones 189, 191

nucleus accumbens and motor activity 535 nucleus of Darkschewitsch cerebellar input 143 nucleus of the accessory olfactory tract accessory olfactory input 539 nucleus of the diagonal band projection to olfactory bulb 541 projection to piriform cortex 544 nucleus of the lateral olfactory tract 519 nucleus prepositus hypoglossi cholinergic mossy fibers 115, 285, 287, 304 corticotropin-releasing factor in mossy fibers 287 GABAergic projection to inferior olive 157, 237 nucleus reticularis tegmenti pontis mossy fibers 165, 291 oculomotor nucleus 145 odor preference 548 olfactory bulb see also main and accessory olfactory bulb function 469 olfactory code 469, 472 olfactory epithelium cell types 470 olfactory marker protein

(OMP) in olfactory receptor cells 472, 475 olfactory nerve function 470 juxtaglomerular dopamine modulation 494 olfactory receptor neurons location 469 shape 470 turnover 498 olfactory system and memory 536 and motor activity 534 inputs to striatal patch/ matrix 438 olfactory tubercle

581

Subject

index

olfactory input 518 olivocerebellar projection see inferior olive opiate receptors in striatum 450 orbitofrontal cortex 533 orphan glutamate receptors cerebellum 72 ovarian cancer cerebellum 48

P400 protein in Purkinje cells 27 paleocortex 519 parafascicular nucleus input to striatum 435 paraflocculus, cerebellum 136 paramedian pontine reticular formation mossy fibers 291 paramedian reticular nucleus mossy fibers 291 paramedian sulcus, cerebellum 172 paraneoplastic diseases cerebellar immunoreactivity 47 parasolitary nucleus GABA projection to inferior olive 234, 237 Parkinson's disease basal ganglia 371,453 olfactory deficits 473 parvalbumin anterior olfactory nucleus 514 climbing fibers 269, 275 inferior olive 282 in some nigral GABA neurons 456 mossy fibers 285 Purkinje cells 36, 37, 220 stellate/basket cells 85 patch/matrix see striatum patchy mosaic, mossy fibers 301 pedunculopontine nucleus to cerebellum 118 PEP-19 polypeptide in Purkinje cells 38 peri-allocortex input to striatal patch/ matrix 438

582

periamygdaloid cortex 522 periaqueductal gray input from central amygdalar nucleus 534 perihypoglossal nuclei mossy fibers 285 pes pontis 291 phospholipase C in Purkinje cells 25, 32 piriform cortex architecture 519 cholinergic actions 545 cholinergic inputs 544 cholinergic receptors 544 commissural and associational connections 527, 527 dopamine innervation 553 glutamate receptors 507 glutamatergic projections 525 interneurons 522, 523 intrinsic connections 526 main olfactory bulb input 524 mitral cell input 491 noradrenergic input 548 norepinephrine actions 550 projection to hypothalamus 528 projection to olfactory bulb 524 projection to striatal patch/ matrix 438 pyramidal cells 521,522 seizurogenic focus 529 serotonin actions 552 serotoninergic inputs 551 transmitter receptors 529 pontine gray see basal pontine gray pontine nuclei see basal pontine gray pontocerebellar projection 291,293, 295 posterior amygdalar nucleus connections 539 posterolateral fissure, cerebellum 138 P-path antigen parasagittal zones 189 precerebellar nuclei peptidergic mossy

fibers 305 prelimbic cortex input to striatal patch/ matrix 432 preoptic region (area) 539 protein kinase C juxtaglomerular cells 486 parasagittal zones 193 Purkinje cells 32, 33 protoplasmic islands of Held 4 Purkinje cell see cerebellar cortex Purkinje cell specific glycoprotein in development 220 pyramis, cerebellum 135 quinpirole striatal changes 448 red nucleus cerebellar projection to 143 projection to cerebellar nuclei 303 projection to inferior olive 235, 261 restiform body 284 reticular formation serotoninergic input to cerebellum 103 reticular tegmental nucleus see nucleus reticularis tegmenti pontis rubrospinal tract to inferior olive 235

schizophrenia olfactory deficits 473 secretogranin unipolar brush cells 89 semicircular fibers, cerebellum 284 serotonin (5-HT) input to cerebellum 103, 167, 303 input to inferior olive 231, 239 input to olfactory bulb 550 serotonin receptors cerebellum 104 olfactory bulb 551 piriform cortex 552 sexual behavior

Subject index and accessory olfactory input 539 sexual dimorphism accessory olfactory system 541 SKF38393 striatal changes 448 social odors 540 somatostatin climbing fibers 275 fastigial nucleus 164 Golgi cells 89 juxtaglomerular cells 481 Purkinje cells 24 somesthetic corpus cerebelli (Larsell) calretinin staining 207 definition 207 spinal cord collaterals to cerebellar nuclei 302 mossy fibers 165, 287 zonal arrangement of mossy fibers 293, 297 spinal trigeminal nucleus mossy fibers 301 spinocerebellar tracts 287, 289, 293 spiny projection neuron striatum 377 stem cells olfactory epithelium 472 striatum cellular interactions 451 cortical input 377, 413 ff. D2 dopamine receptors 447 dopamine receptor-mediated peptide regulation 447 dual striatal/nigral outputs 421 interneurons 377, 390, 452 cholinergic interneuron 391,394 cortical inputs 394 GABAergic interneuron 392 giant K611iker interneuron 391 inputs 394 small aspiny interneutons 391

spidery neuron 391 large aspiny neurons 377 medium aspiny neurons 377 medium spiny neuron see striatum, spiny projection neuron multiple discontinuous cortical inputs 417 multiple overlapping cortical inputs 415 output 418 patch matrix compartments cortical inputs 431,437 general organization 435 matrix to nigral GABA neurons 429 midbrain dopamine 427, 429 patch to dopamine neurons 429 striatal outputs 429 regional differences differential dopamine inputs 456 subdivisions 455 spiny projection neuron (medium-sized) 380 cholinergic input 389 cortical input 382 dopamine receptors 447 GAD content 444 local collateral input 388 local collaterals 452 nigrostriatal input 386 pallidal GABA input 390 striatal GABA interneuron input 389 striatal somatostatin interneuron input 389 subthalamic nucleus input 390 thalamic input 382 striatonigral neuron definition 443 dynorphin content 445 GAD content 433 substance P content 445 striatopallidal neuron definition 443 enkephalin content 445

GAD content 433 submedial nucleus (thalamus) olfactory input 529, 533 substance P accessory olfactory granule cells 538 D2 receptor modulation of in striatum 447 juxtaglomerular cells 481 substance P receptor parasagittal zones 189 substantia nigra compacta inputs 404 dopaminergic neurons 402 GABA neurons 402, 535 neuron morphology 402 reticulata output subregions 423 reticulata output to superior colliculus 424 reticulata pallidal input 403 reticulata striatal input 403 reticulata subthalamic input 403 reticulata thalamic projection 407 subthalamic nucleus cortical inputs 400 globus pallidus input 400 lesions 453 neurons (glutamatergic) 400 output 402 superior cerebellar peduncle location 143 superior colliculus 301 superior vestibular nucleus 143 sustentacular cells olfactory epithelium 470 taenia tecta 516 taurine in Purkinje cells 21 parasagittal zones 190 T-cell antigens in Purkinje cells 39 thalamus inputs to striatum 435 thyrotropin-releasing hormone juxtaglomerular neurons 486 Tourette's syndrome

583

Subject index basal ganglia 371,453 trigeminocerebeUar projection 301 tuberomammillary nucleus histamine projection to cerebellum 111 tyrosine hydroxylase Purkinje cells 24 uncinate tract from fastigial nucleus 143 vasoactive intestinal polypeptide juxtaglomerular cells 481 vasopressin receptor parasagittal zones 189 ventral lateral medulla 534 ventral pallidum 535

584

ventral striatum and nucleus accumbens 456 and olfactory tubercle 518 ventral tegmental area projection to cerebellar nuclei 168 vermis caudal 211, 215 vestibular nuclei mossy fibers to cerebellum 285, 293 parvalbumin in mossy fibers 285 projection to inferior olive 157, 234, 237 zebrin input 203 vestibulocerebellum (Larsell) 207

vibrissal receptive fields, cerebellum 297 vimentin Bergmann glial cells 130 visinine in Purkinje cells 39 vomeronasal organ and behavior 539 physiology 473 zebrins in Purkinje cells 39, 41, 164, 175 parasagittal zones 189, 201, 297 projections to cerebellar nuclei 203